UNIVERSIDADE ESTADUAL DE CAMPINAS
FACULDADE DE ENGENHARIA DE ALIMENTOS
DEPARTAMENTO DE ENGENHARIA DE ALIMENTOS
EXTRAÇÃO, MICRONIZAÇÃO E ESTABILIZAÇÃO DE PIGMENTOS
FUNCIONAIS: CONSTRUÇÃO DE UMA UNIDADE MULTIPROPÓSITO
PARA DESENVOLVIMENTO DE PROCESSOS COM FLUIDOS
PRESSURIZADOS
Diego Tresinari dos Santos
Orientador: Profa. Dra. Maria Angela de Almeida
Meireles.
Tese apresentada à Faculdade de Engenharia de
Alimentos da Universidade Estadual de Campinas,
para Obtenção do Título de Doutor em Engenharia
de Alimentos.
CAMPINAS-SP
Fevereiro de 2011
FICHA CATALOGRÁFICA ELABORADA PELA
BIBLIOTECA DA FEA – UNICAMP
Sa59e
Santos, Diego Tresinari dos
Extração, micronização e estabilização de pigmentos funcionais:
construção de uma unidade multipropósito para desenvolvimento de
processos com fluidos pressurizados / Diego Tresinari dos Santos. -Campinas, SP: [s.n], 2011.
Orientador: Maria Angela de Almeida Meireles
Tese (doutorado) – Universidade Estadual de Campinas. Faculdade
de Engenharia de Alimentos.
1. Estabilização. 2. Extração. 3. Fluidos Pressurizados. 4.
Micronização. 5. Pigmentos funcionais. I. Meireles, Maria Angela
de Almeida. II. Universidade Estadual de Campinas. Faculdade de
Engenharia de Alimentos. III. Título.
cars/bibfea
Título em inglês: Extraction, micronization and stabilization of functional pigments:
construction of multipurpose unit for pressurized fluid process
development
Palavras-chave em inglês (Keywords): Stabilization, Extraction, Pressurized Fluids,
Micronization, Functional pigments
Titulação: Doutor em Engenharia de Alimentos
Banca examinadora: Maria Angela de Almeida Meireles
Silvânia Regina Mendes Moreschi
Carlos Raimundo Ferreira Grosso
Reinaldo Camino Bazito
Juliana Martin do Prado
Data da Defesa: 22/02/2011
Programa de Pós Graduação em Engenharia de Alimentos
ii
Este exemplar corresponde à redação final da tese defendida em 22/02/2011 por Diego
Tresinari dos Santos aprovado pela comissão julgadora em 22/02/2011.
________________________________________________
Profa. Dra. Maria Angela de Almeida Meireles
FEA / UNICAMP
Orientador
________________________________________________
Profa. Dra. Silvânia Regina Mendes Moreschi
Universidade Tecnológica Federal do Paraná
Membro
________________________________________________
Prof. Dr. Carlos Raimundo Ferreira Grosso
FEA / UNICAMP
Membro
________________________________________________
Prof. Dr. Reinaldo Camino Bazito
IQ / USP
Membro
________________________________________________
Dra. Juliana Martin do Prado
FEA / UNICAMP
Membro
________________________________________________
Dra. Carmen Lucia Queiroga
CPQBA / UNICAMP
Suplente
________________________________________________
Dr. Flávio Cortiñas Albuquerque
CENPES / PETROBRÁS
Suplente
________________________________________________
Profa. Dra. Miriam Dupas Hubinger
FEA / UNICAMP
Suplente
iii
Todas as inovações eficazes são surpreendentemente simples.
Na verdade, o maior elogio que uma inovação pode receber é haver quem diga:
isto é óbvio. Por que não pensei nisso antes?
Peter Drucker
iv
AGRADECIMENTOS
Á Deus, por tudo que me há propiciado seja profissionalmente ou pessoalmente.
À UNICAMP, por toda a sua estrutura.
Ao CNPq, pela concessão da bolsa e financiamento deste projeto de pesquisa.
À CAPES, pelo auxílio financeiro (PROEX) para participação no 10 º Congresso Brasileiro
de Polímeros em Foz do Iguaçu-PR.
À Profa. Dra. Maria Angela de Almeida Meireles, pela brilhante orientação, confiança,
entusiasmo e exemplo como orientadora, pesquisadora e professora, bem como aos
ensinamentos de não mensurável valor transmitidos a mim de ordem profissional e pessoal.
Ao Pesquisador Ariovaldo Astini, pela amizade e valiosa contribuição de ordem técnica e
intelectual durante o desenvolvimento deste projeto de pesquisa.
Aos Pesquisadores Prof. Dr. Elton Franceschi, Prof. Dr. Paulo de Tarso Vieira e Rosa e aos
membros da banca examinadora, pelas sugestões e ensinamentos fundamentais para a
realização deste trabalho.
Às Pesquisadoras Profa. Dra. Marisa Beppu e Dra. Carmem Queiroga, por disponibilizarem
seus laboratórios para a realização de alguns experimentos referentes a este projeto de
pesquisa.
À minha amiga, parceira de dança, investidora, “marida” e Pesquisadora Ms. Juliana
Queiroz Albarelli, pelas inúmeras “trocas de idéias” sobre o desenvolvimento desta tese e de
outros trabalhos que desenvolvemos em parceria, bem como por ter contribuído
v
significativamente no desenvolvimento dos experimentos relacionados à estabilização dos
extratos antociânicos.
Aos colegas de trabalho Priscilla Veggi, Rodrigo Cavalcanti, Helmut Navarro e Carolina
Albuquerque, pelo ânimo e disponibilidade de desenvolvermos alguns trabalhos em
paralelo que resultaram e ainda resultarão em algumas publicações.
Aos Pesquisadores e amigos Prof. Dr. Silvio Silvério da Silva, Prof. Dr. Attílio Converti,
Prof. Dr. Victor Haber Pérez, Dr. Boutros Fouad Sarrouh, Profa. Dra. Lilian Masson, Prof.
Dr. Michael Oelgemöller e Profa. Dra. Maria José Cocero, que com seus valiosos
ensinamentos e conselhos contribuíram para a construção do profissional que me tornei.
À minha Mãe, por ter sempre se preocupado prioritariamente com a minha educação e
formação, as quais foram os alicerces para a chegada no patamar que estou, bem como pelo
amor incondicional.
Ao meu Pai, à minha Vó, Tio Fernando, Tia Maria, Tia Flor e a todos os familiares que,
mesmo de longe, participaram me apoiando e incentivando em todos os momentos.
Aos amigos e colegas do DEA pelas horas agradáveis que compartilhamos.
vi
ÍNDICE
RESUMO........................................................................................................................ XVII
ABSTRACT ..................................................................................................................... XIX CAPÍTULO 1 - INTRODUÇÃO E OBJETIVOS ............................................................. 1 1.1 Introdução ....................................................................................................................... 1 1.2 Objetivos da Pesquisa..................................................................................................... 3 1.2.1 Geral ...................................................................................................................... 3 1.2.2 Específicos ............................................................................................................. 3 1.3 Estrutura da Tese de Doutorado ................................................................................... 4 CAPÍTULO 2 - REVISÃO BIBLIOGRÁFICA................................................................. 9 2.1. Pigmentos Funcionais .................................................................................................... 9 2.1.1 Flavonóides ......................................................................................................... 10 2.1.2 Carotenóides........................................................................................................ 13 2.2. Métodos de Extração ................................................................................................... 15 2.2.1 Extração com Fluidos Pressurizados .................................................................. 17 2.2.1.1 Extração com Líquidos Pressurizados .......................................................... 18 2.2.1.2 Extração com CO2 supercrítico .................................................................... 19 2.2.2 Métodos de Extração Assistida ........................................................................... 20 2.2.2.1. Com ultrassom ............................................................................................. 20 2.2.2.2. Com CO2 a alta pressão ............................................................................... 21 2.3. Formação de Partículas .............................................................................................. 22 2.4.1 Para Extração ..................................................................................................... 23 2.4.1.1 Para Extração com Líquidos Pressurizados .................................................. 25 2.4.1.2 Para Extração com CO2 Supercrítico........................................................... 25 2.4.1.3 Para Extração assistida com ultrassom ......................................................... 26 2.4.1.4 Para Extração assistida com CO2 a alta pressão ........................................... 27 2.4.2 Para Formação de Partículas ............................................................................. 28 2.4.2.1 Para Formação de Partículas Via SAS ......................................................... 28 2.4.2.2 Para Formação de Partículas Via RESS ....................................................... 29 Referências .......................................................................................................................... 29 CAPÍTULO 3 - DETALHAMENTO DA CONSTRUÇÃO E FUNCIONAMENTO DA
UNIDADE MULTIPROPÓSITO ..................................................................................... 37 CAPÍTULO 4 - ANTIOXIDANT PIGMENT EXTRACTION USING A HOMEMADE PRESSURIZED SOLVENT EXTRACTION SYSTEM ................................... 43 Key words ............................................................................................................................ 44 Abstract ............................................................................................................................... 44 vii
4.1 Introduction .................................................................................................................. 44 4.2 Materials and methods ................................................................................................. 47 4.2.1 Plant Material ..................................................................................................... 47 4.2.1 Annatto seeds................................................................................................... 47 4.2.2 Jabuticaba skins ............................................................................................... 47 4.2.2 Extraction Procedures ......................................................................................... 48 4.2.3 Extract Characterization ..................................................................................... 51 4.2.3.1 From Jabuticaba skins .................................................................................. 51 4.2.3.1.1 Anthocyanin content .............................................................................. 51 4.2.3.1.2 Thin-Layer Chromatography (TLC)...................................................... 52 4.2.4 Statistical Analysis............................................................................................... 52 4.3 Results and discussion .................................................................................................. 53 4.3.1 Obtaining Annatto seed extracts ......................................................................... 53 4.3.2 Obtaining Jabuticaba skin extracts ..................................................................... 56 4.4 Conclusions ................................................................................................................... 59 Acknowledgements ............................................................................................................. 59 References ........................................................................................................................... 59 CAPÍTULO 5 - PRESSURIZED LIQUID EXTRACTION OF PHENOLIC
COMPOUNDS FROM JABUTICABA SKINS: OPTIMIZATION STUDY ............... 61 Key words ............................................................................................................................ 62 Abstract ............................................................................................................................... 62 5.1 Introduction .................................................................................................................. 62 5.2 Material and methods .................................................................................................. 65 5.2.1 Plant Material ..................................................................................................... 65 5.2.2 Extraction Procedures ......................................................................................... 65 5.2.3 Extract Characterization ..................................................................................... 67 5.2.4 Statistical Analysis............................................................................................... 69 5.3 Results and discussion .................................................................................................. 70 5.3.1 Effects of process variables on the extraction yield ............................................ 70 5.3.2 Effects of process variables on the recovery of anthocyanins ............................. 71 5.3.3 Effects of process variables on the recovery of phenolic compounds ................. 74 5.3.4 Optimization of the extraction process ................................................................ 77 5.4 Conclusions ................................................................................................................... 82 Acknowledgements ............................................................................................................. 83 References ........................................................................................................................... 83 viii
CAPÍTULO 6 - OPTIMIZATION OF BIOACTIVE COMPOUNDS EXTRACTION
FROM JABUTICABA (MYRCIARIA CAULIFLORA) SKINS ASSISTED BY HIGH
PRESSURE CO2 ................................................................................................................. 87 Key words ............................................................................................................................ 88 Abstract ............................................................................................................................... 88 6.1 Introduction .................................................................................................................. 89 6.2 Material and methods .................................................................................................. 91 6.2.1 Plant material ...................................................................................................... 91 6.2.2 High Pressure Carbon Dioxide Assisted-Extraction (HPCDAE) system............ 91 6.2.3 Extraction Procedures ......................................................................................... 92 6.2.4 Extract Characterization ..................................................................................... 95 6.2.5 Statistical analysis ............................................................................................... 97 6.2.6 Determination of experimental extraction kinetics curves and parameters ........ 97 6.3 Results and discussion .................................................................................................. 97 6.3.1 Effects of process variables on recovery of anthocyanins................................... 97 6.3.2 Effects of process variables on recovery of phenolic compounds ..................... 101 6.3.3 Optimization of the extraction process .............................................................. 105 6.3.4 Experimental extraction kinetics curves using optimum conditions ................. 108 6.4 Conclusions ................................................................................................................. 112 Acknowledgements ........................................................................................................... 114 References ......................................................................................................................... 114 CAPÍTULO 7 - MICRONIZATION AND ENCAPSULATION OF FUNCTIONAL
PIGMENTS USING SUPERCRITICAL CARBON DIOXIDE .................................. 117 Key words .......................................................................................................................... 118 Abstract ............................................................................................................................. 118 7.1 Introduction ................................................................................................................ 119 7.2 Materials and methods ............................................................................................... 121 7.2.1 Materials............................................................................................................ 121 7.2.2 Micronization process via SAS .......................................................................... 124 7.2.3 Encapsulation process via SAS ......................................................................... 127 7.2.4 Encapsulation process via RESS ....................................................................... 128 7.2.5 Characterization and Analysis .......................................................................... 130 7.2.5.1.1 Determination of Precipitation Yield - PY (%) .................................. 131 7.2.5.1.2 Determination of Encapsulation Efficiency (EE (%)) ......................... 131 7.3 Results and discussion ................................................................................................ 132 7.3.1 Micronization process via SAS .......................................................................... 132 7.3.2 Encapsulation process via SAS ......................................................................... 138 ix
7.3.3 Encapsulation process via RESS ....................................................................... 142 7.4 Conclusions ................................................................................................................. 145 Acknowledgements ........................................................................................................... 147 References ......................................................................................................................... 147 CAPÍTULO 8 - STABILIZATION OF ANTHOCYANIN EXTRACT FROM
JABUTICABA SKINS BY ENCAPSULATION USING SUPERCRITICAL CO2 AS
SOLVENT ......................................................................................................................... 153 Key words .......................................................................................................................... 154 Abstract ............................................................................................................................. 154 8.1 Introduction ................................................................................................................ 154 8.2.2 Anthocyanin Extraction ..................................................................................... 156 8.2.3 Encapsulation of Anthocyanin Extract .............................................................. 156 8.2.3.1 By Rapid Expansion of Supercritical Solution (RESS) process ................ 156 8.2.3.1.1 Evaluation of the antioxidant activity of the encapsulated anthocyanin
extract after RESS process ..................................................................................... 161 8.2.3.2 By conventional method ............................................................................. 161 8.2.4 Anthocyanin Stabilization Studies ..................................................................... 162 8.2.4.1 Free and encapsulated extract degradation studies ..................................... 162 8.2.4.2 Thermal analysis of free and encapsulated extracts ................................... 164 8.2.5 Anthocyanin Extract Release Studies ................................................................ 164 8.2.6 Statistical Analysis............................................................................................. 164 8.3 Results and discussion ................................................................................................ 165 8.3.1 Encapsulation of anthocyanin extract by RESS process ................................... 165 8.3.2 Encapsulation of anthocyanin extract by conventional method ....................... 169 8.3.3 Anthocyanin stabilization by encapsulation ...................................................... 170 8.3.4 Release studies................................................................................................... 173 8.4 Conclusions ................................................................................................................. 175 Acknowledgements ........................................................................................................... 176 References ......................................................................................................................... 176 CAPÍTULO 9 - CONCLUSÕES GERAIS ..................................................................... 181 SUGESTÕES PARA TRABALHOS FUTUROS .......................................................... 185 MEMÓRIA DO PERÍODO DO DOUTORADO .......................................................... 187 PRODUÇÃO BIBLIOGRÁFICA ................................................................................... 189 APÊNDICE I - ARTIGO DE REVISÃO - JABUTICABA AS A SOURCE OF
FUNCTIONAL PIGMENTS ........................................................................................... 193 x
APÊNDICE II – DETALHAMENTO DO PROCEDIMENTO DE CÁLCULO DA
CONCENTRAÇÃO DE ANTOCIANINAS E FENÓIS ............................................... 201
APÊNDICE III - ARTIGO DE REVISÃO - CAROTENOID PIGMENTS
ENCAPSULATION: FUNDAMENTALS, TECHNIQUES AND RECENT TRENDS
............................................................................................................................................ 207
APÊNDICE IV - MANUAL DE OPERAÇÃO DA UNIDADE MULTIPROPÓSITO
............................................................................................................................................ 219
LISTA DE TABELAS
Tabela 2.2.1.1 - Constantes críticas de alguns fluidos com interesse em extração (MENDES
et al., 2003) ........................................................................................................................... 17
Table 4.3.1.1 - Percentage of Residual Extract (%) deposited in the exit tubing line when
using different SFE systems/configurations. ........................................................................ 55
Table 5.2.2.1 - The experimental design of phenolic compound extraction from jabuticaba
skins ...................................................................................................................................... 66
Table 5.3.4.1 - Optimum PLE conditions for the extraction yields, anthocyanins and total
phenols from jabuticaba skins .............................................................................................. 77
Table 5.3.4.2 - Predicted and experimental values of responses under optimum PLE
conditions (50 bar, temperature of 80 °C and static extraction time of 9 min) and
experimental values responses obtained by Conventional Low Pressure Liquid Extraction
(LPLE) .................................................................................................................................. 79
Table 6.3.1.1 - 23 full factorial design for HPCD Assisted-Extraction from jabuticaba skins
and the total anthocyanin and phenolic contents of the extracts .......................................... 99
Table 6.3.1.2 - Regression coefficients of the model for the response variables ............... 101
Table 6.3.3.1 - Predicted and experimental values of response variables under optimum
HPCDAE conditions (at 117 bar, temperature of 80 °C, of 20 % and 20 minutes of
extraction) and experimental values of response variables obtained by control HPCDAE
experiment (at atmospheric pressure, temperature of 80 ºC and 20 minutes of extraction)
and by PLE experiment (at 117 bar, temperature of 80 ºC, 20 minutes of extraction and
flow rate of 1 mL of acidified water/min) .......................................................................... 108
Table 6.3.4.1 - Experimental values of the steady-state extraction Y*, the time t* to reach
the Y* and the mass transfer rate M* for the recovery of anthocyanins and phenolic
compounds .......................................................................................................................... 111
LISTA DE FIGURAS
Figura 2.1.1.1 - Estrutura base dos flavonóides. .................................................................. 11
Figura 2.1.1.2 - Cascas de jabuticaba. .................................................................................. 12
xi
Figura 2.1.2.1 - Estrutura do β-caroteno............................................................................... 13
Figura 2.1.2.2 - Sementes de urucum. .................................................................................. 15
Figura 2.2.1.2.1 - Diagrama de fase pressão-temperatura do dióxido de carbono ............... 20
Figura 2.4.1.1.1 - Equipamento comercial para extração com líquidos pressurizados. a)
ASE® 150; b) ASE® 350 (DIONEX, 2010).......................................................................... 24
Figura 2.4.1.1.2 - Equipamento comercial para extração com líquidos pressurizados
(FLUID MANAGEMENT SYSTEMS, 2010) ..................................................................... 24
Figura 2.4.1.2.1 - Equipamento comercial para extração com CO2 supercrítico (APPLIED
SEPARATIONS, 2010) ........................................................................................................ 26
Figura 2.4.1.3.1 - Equipamento comercial para extração assistida com ultrassom
(HIELSCHER, 2010) ........................................................................................................... 27
Figura 2.4.2.1.1 - Equipamento comercial para formação de partículas via SAS (THAR
TECHNOLOGIES, 2010) .................................................................................................... 28
Figura 2.4.2.2.1 - Fluxograma da unidade experimental comercial da Thar Technologies
RESS-100. 1 - Reator; 2 - Agitador; 3 – Termostato; 4, 7 e 8 - válvulas; 5 -Medidor de
fluxo; 6 - Bomba de alta pressão; 9 - Dispositivo de expansão; 10 - Trocador de calor para
aquecimento; 11 - Câmara de expansão; 12 - Trocador de calor para resfriamento; 13 Computador (GIL’MUTDINOV et al., 2009). ..................................................................... 29
Figura 3.1 - Foto da parte estrutural da unidade. .................................................................. 38
Figura 3.2 - Fotos da unidade multipropósito. A - Vista de frente da unidade; B - Sistema
ultrassônico quando acoplado à unidade; C - Vista do sistema de aquecimento do vasos de
pressão menor: D - Vista lateral da unidade. ........................................................................ 40
Figura 3.3 - Foto da serpentina imersa no banho termostatizado de resfriamento. .............. 42
Figure 4.2.2.1 - Schematic diagram of the home-made pressurized solvent extraction
system. 1 CO2 cylinder; 2 CO2 filter; 3 Manometers; 4 Valves; 5 Thermostatic bath; 6 CO2
Pump; 7 Back pressure regulator; 8 HPLC pump; 9 Solvent resevoir; 10 Extraction cell; 11
Micrometric valve with a heating system; 12 Temperature controller; 13 Sampling bottle.48
Figure 4.2.2.2 - Schematic diagram of the home-made pressurized solvent extraction
system. 1 CO2 cylinder; 2 CO2 filter; 3 Manometers; 4 Valves; 5 Thermostatic bath; 6 CO2
Pump; 7 Back pressure regulator; 8 HPLC pump; 9 Solvent reservoir; 10 Extraction cell; 11
Ultrasound bath; 12 Heating bath; 13 Micrometric valve with a heating system; 14
Temperature controller; 15 Sampling bottle......................................................................... 50
Figure 4.3.1.1 - Recovery of pigments from Annatto seeds as a function of time using
different SFE systems/configurations: ■ Commercial SFE; □ Home-made SFE using
electric heating; ● Home-made Ultrasound Assisted SFE; ○ Home-made SFE using water
bath heating. ......................................................................................................................... 54
Figure 4.3.1.2 - Extraction yields (%) (after 240 min) using different SFE
systems/configurations. ........................................................................................................ 55
xii
Figure 4.3.2.1 - Thin-layer chromatography (TLC) plates (1 - Extract obtained at 30
MPa/333 K run 1; 2 – Extract obtained at 30 MPa/333 K run 2; a) Revealed using DPPH
reagent on visible light; b) Revealed using Natural products (NP) reagent on light (365 nm);
c) Revealed using anisaldehide-sulphuric acid reagent on visible light; d) Revealed using
anisaldehide-sulphuric acid reagent on ultraviolet light (365 nm). ...................................... 58
Figure 5.2.2.1 - Pressurized liquid extraction set-up. ........................................................... 66
Figure 5.3.1 - Three-dimensional response surfaces of the influence of extraction
temperature and static extraction time on the extraction yield. ............................................ 70
Figure 5.3.2.1 - Effect (p<0.05) of extraction variables on the recovery of anthocyanins: a)
at 50-100 bar, 40-80 ºC and 3-9 min; b) at 80-120 ºC and 9-15 min at a fixed extraction
pressure of 50 bar (1, pressure; 2, temperature; 3, static time)............................................. 72
Figure 5.3.2.2 - Three-dimensional response surfaces of the influence of temperature and
static extraction time on recovery of anthocyanins .............................................................. 73
Figure 5.3.3.1 - Effect (p<0.05) of extraction variables on the recovery of total phenolic
compounds: a) at 50-100 bar, 40-80 ºC and 3-9 min; b) at 80-120 ºC and 9-15 min at a
fixed extraction pressure of 50 bar (1, pressure; 2, temperature; 3, static time). ................. 75
Figure 5.3.3.2 - Three-dimensional response surfaces of the influence of temperature and
static extraction time on recovery of total phenolic compounds. ......................................... 76
Figure 5.3.4.1 - Kinetics curves: a) overall extraction yield; b) recovery of anthocyanins; c)
recovery of total phenolic compounds under optimum PLE conditions (50 bar, temperature
of 80 °C and 9 min of static extraction time). ...................................................................... 81
Figure 6.2.1 - Diagram of High Pressure Carbon Dioxide Assisted-Extraction (HPCDAE)
system. 1 CO2 cylinder; 2 CO2 filter; 3 Manometers; 4 Valves; 5 Thermostatic bath; 6
Pump; 7 Back pressure regulator; 8 Heating bath; 9 High pressure vessel; 10
Thermocouple; 11 Temperature controllers; 12 Micrometric valve with a heating system. 92
Figure 6.3.1.1 - Three-dimensional response surfaces of the influence of extraction pressure
and temperature on recovery of anthocyanins. ................................................................... 100
Figure 6.3.1.2 - Two-dimensional response surfaces of the influence of extraction pressure
and temperature on recovery of anthocyanins. ................................................................... 100
Figure 6.3.2.1 - Three-dimensional response surfaces of the influence of extraction pressure
and temperature on the recovery of phenolic compounds. ................................................. 103
Figure 6.3.2.2 - Two-dimensional response surfaces of the influence of extraction pressure
and temperature on the recovery of phenolic compounds. ................................................. 104
Figure 6.3.3.1 - Three-dimensional response surfaces of the influence of the extraction
variables on the desirability. ............................................................................................... 106
Figure 6.3.3.2 - Profiles of the predicted values and desirability of the extraction variables.
............................................................................................................................................ 106
xiii
Figure 6.3.4.1 - Kinetics curves for the recovery of anthocyanins a) under optimum
HPCDAE conditions (117 bar, temperature of 80 °C and of 20 %; b) obtained by control
HPCDAE experiment (at atmospheric pressure, temperature of 80 ºC and 20 minutes of
extraction) and c) PLE experiment (at 117 bar, temperature of 80 ºC, 20 minutes of
extraction and flow rate of 1 mL/min of acidified water/min). .......................................... 109
Figure 6.3.4.2 - Kinetics curves for the recovery of phenolic compounds a) under optimum
HPCDAE conditions (117 bar, temperature of 80 °C and of 20 %; b) obtained by control
HPCDAE experiment (at atmospheric pressure, temperature of 80 ºC and 20 minutes of
extraction) and c) PLE experiment (at 117 bar, temperature of 80 ºC, 20 minutes of
extraction and flow rate of 1 mL/min of acidified water/min). .......................................... 110
Figure 7.2.1.1 - SEM micrograph of unprocessed quercetin sample. ................................ 122
Figure 7.2.1.2 - SEM micrograph of unprocessed β-carotene sample. .............................. 122
Figure 7.2.1.3 - SEM micrograph of unprocessed rutin sample. ........................................ 124
Figure 7.2.2.1 - Schematic diagram of the SAS apparatus. 1 CO2 cylinder; 2 CO2 filter; 3
manometers; 4 valves; 5 thermostatic bath; 6 CO2 pump; 7 back pressure regulator; 8
solution reservoir; 9 solution pump; 10 thermocouple; 11 precipitation vessel; 12 heating
bath; 13 temperature controllers; 14 micrometric valve with a heating system; 15 glass
flask; 16 glass float rotameter; 17 flow totalizer. ............................................................... 126
Figure 7.2.4.1 - Schematic diagram of the RESS apparatus. 1 CO2 cylinder; 2 CO2 filter; 3
manometers; 4 valves; 5 thermostatic bath; 6 CO2 pump; 7 back pressure regulator; 8 preexpansion vessel; 9 micrometric valve with a heating system; 10 temperature controller; 11
nozzle; 12 expansion vessel................................................................................................ 129
Figure 7.3.1.1.1 - SEM micrograph of quercetin micronized particles obtained by SAS
process. ............................................................................................................................... 133
Figure 7.3.1.1.2 - SEM micrograph of quercetin micronized particles obtained by
conventional process........................................................................................................... 133
Figure 7.3.1.1.3 - Size distribution of quercetin particles obtained by SAS and conventional
processes. ............................................................................................................................ 135
Figure 7.3.1.2.1 - SEM micrograph of β-carotene micronized particles obtained by SAS
process. ............................................................................................................................... 136
Figure 7.3.2.1.1 - Optical micrographs of: 1) PEG sample; 2) bixin-rich extract
encapsulated in PEG by SAS process (CO2 flow rate of 0.6 kg.h-1; mass ratio between core
material and PEG of 1:10); 3) rutin encapsulated in PEG by RESS process (mass ratio
between core material and PEG of 1:10); 4) anthocyanin-rich extract encapsulated in PEG
by RESS process (mass ratio between core material and PEG of 1:10)............................. 140
Figure 7.3.2.1.2 - SEM micrographs of: 1) bixin-rich extract encapsulated in PEG by SAS
process (CO2 flow rate of 0.6 kg.h-1; mass ratio between core material and PEG of 1:10);
2) rutin encapsulated in PEG by RESS process (mass ratio between core material and PEG
of 1:2); 3) rutin encapsulated in PEG by RESS process (mass ratio between core material
xiv
and PEG of 1:10); 4) anthocyanin-rich extract encapsulated in PEG by RESS process (mass
ratio between core material and PEG of 1:10). .................................................................. 141
Figure 8.8.2.3.1.1 - Schematic diagram of the RESS apparatus. 1 CO2 cylinder; 2 CO2
filter; 3 manometers; 4 valves; 5 thermostatic bath; 6 CO2 pump; 7 back pressure regulator;
8 pre-expansion vessel; 9 micrometric valve with a heating system; 10 temperature
controller; 11 nozzle; 12 expansion vessel. ........................................................................ 158
Figure 8.2.3.2.1 - Encapsulation by entrapment in Ca-alginate beads. .............................. 162
Figure 8.3.1.1 - Optical micrographs of unprocessed PEG (a), Encapsulated anthocyanin
extract – T: 313.15 K; P: 100 bar (b), Encapsulated anthocyanin extract – T: 323.15 K; P:
100 bar (c)........................................................................................................................... 166
Figure 8.3.1.2 - Influence of different proportions of anthocyanin extract:PEG on the
percentage of encapsulated (gray bars); encapsulation efficiency (black bars) ................. 168
Figure 8.3.1.3 - Antioxidant activity of encapsulated anthocyanin extracts obtained using
different operating RESS conditions (gray symbols), the anthocyanin extract (♦) and pure
synthetic BHT (■); without any antioxidant compound (▲) ............................................. 169
Figure 8.3.3.1 - Degradation of free anthocyanin extract at different environments. ........ 170
Figure 8.3.3.2 - Degradation of encapsulated anthocyanin extract at different environments
by RESS process and conventional method. ...................................................................... 171
Figure 8.3.4.3 - DSC thermograms of free anthocyanin extract, Ca-alginate beads, PEG,
anthocyanin extract encapsulated in Ca-alginate beads and anthocyanin extract
encapsulated in PEG. .......................................................................................................... 173
Figure 8.3.4.1 - The cumulative release of anthocyanin extract from the encapsulated
systems obtained by RESS process and conventional method at pH 1.4 and temperature 37
ºC. ....................................................................................................................................... 174
Figura A1- Esquema das transformações estruturais das antocianinas em função do pH
gerando soluções coloridas. ................................................................................................ 201
Figura A2- Espectro de varredura para os extratos de casca de jabuticaba obtidos utilizando
diferentes métodos de extrações. ........................................................................................ 203
Figura A3 - Curva de calibração previamente construída de ácido gálico (AG) ............... 206
xv
NOMENCLATURAS/ABREVIATURAS
RESS - Rapid Expansion of Supercritical Solutions
SAS - Supercritical fluid Anti-Solvent
SSI - Supercritical Solvent Impregnation
PGSS - Particles from Gas Saturated Solutions
SFEE - Supercritical Fluid Extraction of Emulsions
GRAS - Generally Recognised as Safe
PEG - PolyEthylene Glycol
SFE - Supercritical Fluid Extraction
PLE - Pressurized Liquid Extraction
USFE - Ultrasound-Assisted Supercritical Fluid Extraction
LPLE - Low Pressure Liquid Extraction
ASE – Accelerated Solvent Extraction
PSE - Pressurized Solvent Extraction
RSM - Response Surface Methodology
HPCDAE - High Pressure Carbon Dioxide Assisted Extraction
SCFs - SuperCritical Fluids
xvi
RESUMO
A indústria de alimentos está constantemente à procura de compostos que apresentam
propriedades físicas e químicas para melhorar seus produtos. A maioria destes compostos
são aditivos com propriedades antioxidantes, corantes ou aditivos com efeitos positivos
sobre a saúde humana. Aditivos naturais são sempre preferíveis aos compostos sintéticos.
Flavonóides e carotenóides são duas das principais classes de pigmentos funcionais pelas
quais as indústrias de alimentos, cosmética e farmacêutica têm apresentado maior interesse.
No entanto, estes compostos apresentam uma série de limitações ao serem aplicados em
produtos processados. Diversos fatores, tais como luz, temperatura, pH, entre outros,
desencadeiam a degradação oxidativa destes pigmentos funcionais limitando não só a
aplicação final destes, mas também restringindo toda a cadeia do processo: desde a escolha
do método de extração do pigmento da fonte vegetal até o tratamento que o produto
formulado irá sofrer após a sua formulação passando pela escolha do método de redução do
tamanho e/ou encapsulação das partículas visando a melhora da taxa de dissolução,
biodisponibilidade e estabilidade destes compostos. Tecnologias de extração, micronização
e estabilização de pigmentos funcionais por encapsulação em matrizes poliméricas
utilizando fluidos pressurizados podem representar uma alternativa ambientalmente correta,
uma vez que estão incluídas no conceito de "química verde" e do desenvolvimento
sustentável, e economicamente viável em relação aos respectivos métodos convencionais,
onde grandes quantidades de solventes orgânicos, longos tempos de processo e altas
temperaturas são requeridas, o que pode promover a degradação, isto é, perda de cor e
capacidade antioxidante, condições estas normalmente utilizadas nos processos
convencionais. Adicionalmente, processos de extração e formação de partículas utilizando
fluidos supercríticos permitem um fácil e eficiente controle do processo através de
pequenas variações nas condições de operação (Pressão, Temperatura, etc.). Apesar de
comercialmente encontrarem-se disponíveis equipamentos distintos para cada processo
mencionado uma unidade para pesquisa que possibilite o estudo de diferentes processos
com fluidos pressurizados proporcionaria uma melhor relação custo-benefício associada a
esta tecnologia. Portanto, uma unidade multipropósito para o desenvolvimento de processos
com fluidos pressurizados que possibilite a extração e formação de partículas de pigmentos
xvii
funcionais, bem como de outros compostos bioativos em um único equipamento foi
projetada, construída e testada. Processos de extração utilizando CO2 supercrítico ou
líquidos pressurizados como solventes, assistidos por dióxido de carbono a alta pressão; de
formação de partículas encapsuladas ou não via RESS (Rapid Expansion of Supercritical
Solutions) e SAS (Supercritical fluid Anti-Solvent) foram desenvolvidos na unidade
multipropósito produzindo resultados semelhantes aos obtidos por equipamentos similares,
reprodutíveis e melhores do que quando utilizando processos convencionais.
Palavras-chave: Estabilização, Extração, Fluidos Pressurizados, Micronização, Pigmentos
Funcionais.
xviii
ABSTRACT
The food industry is continuously searching compounds that present physical and chemical
properties to improve their products. Most of them are additives with antioxidant
properties, colorants or additives with positive effects to human health. Natural additives
are always preferred to synthetic compounds. Flavonoids and carotenoids are two of the
major functional pigments class that food, cosmetic and pharmaceutical industries are more
interested recently. Nevertheless, these compounds present a series of limitations when
applied to processed products. Several factors, such as light, temperature, pH, among
others, trigger the oxidative degradation of theses functional pigments limiting not only
their final applications, but also restricting all the process chain: from the choice of the
extraction method of the pigment from the vegetable source, passing through the choice of
the particle reduction and/or encapsulation technique aiming the improvement of the
dissolution rate, biodisponibility and stability of these compounds. Technologies for
extraction, micronization and stabilization of functional pigments into polymeric matrices
using supercritical fluids may represent an environmentally friend alternative, once they are
inserted in the concept of green chemistry and sustainable development, and economically
viable comparing to conventional methods, wherein large amounts of organic solvents, long
process time and high temperatures are required, that can promote degradation, i. e., color
and antioxidant activity loss, conditions normally employed on conventional processes.
Moreover, extraction and particle formation processes utilizing supercritical fluids permit
an easy and efficient process control with little variation on operational conditions
(Pressure, Temperature, etc.). Despite distinct commercial equipments are available to carry
out each mentioned process a unit for research that can be used to carry out different
processes with pressurized fluids would lead to a better cost-benefit relation associated to
this technology. Therefore, a multipurpose unit to develop processes with pressurized fluids
that can be used for extraction and particle formation purposes of functional pigments, as
well as of other bioactive compounds using the same apparatus was designed, constructed
and tested. Extraction processes using supercritical CO2, employing pressurized liquid
solvents, assisted by high pressure carbon dioxide; particle formation processes to obtain
encapsulated or non encapsulated particles via RESS (Rapid Expansion of Supercritical
xix
Solutions) and SAS (Supercritical Anti-Solvent) were done using the multipurpose unit
producing comparable experimental results to those obtained by similar equipments. Good
reproducibility and better results than those obtained using conventional processes were
observed employing our home-made apparatus.
Keywords: Stabilization, Extraction, Pressurized Fluids, Micronization, Functional
Pigments.
xx
CAPÍTULO 1 - INTRODUÇÃO E OBJETIVOS
1.1 Introdução
O aumento da preocupação dos consumidores sobre o uso de aditivos sintéticos
em produtos tem impulsionado as indústrias alimentícias, de cosméticos e farmacêutica em
direção à substituição destes aditivos por produtos naturais. Entretanto, dificuldades têm
sido encontradas devido, principalmente, à instabilidade destes compostos. Um exemplo
são os pigmentos funcionais extraídos de fontes vegetais, tais como as antocianinas e
carotenóides (KONG et al., 2003; MIKI, 1991).
Os flavonóides e os carotenóides são responsáveis por uma grande variedade de
cores presentes em vegetais, flores, frutas e produtos derivados. Oriundos do metabolismo
secundário das plantas, ambas as classes de compostos são de grande importância para a
sobrevivência delas, bem como quando ingeridos são responsáveis por diversos benefícios
à saúde devido às suas atividades biológicas. Entre tais benefícios encontram-se a redução
da incidência de muitas doenças oxidativas, inflamatórias, entre outras (REYNERTSON et
al., 2008; ARTS; HOLLMAN, 2005).
Diversos fatores, tais como luz, temperatura, pH, entre outros, desencadeiam a
degradação oxidativa destes pigmentos funcionais (SANTOS; MEIRELES, 2009, 2010)
limitando não só a aplicação final destes, mas também restringindo toda a cadeia do
processo: desde a escolha do método de extração do pigmento da fonte vegetal até o
tratamento que o produto final irá sofrer após a sua formulação, passando pela escolha do
método de redução do tamanho (micronização) e/ou encapsulação das partículas visando a
melhora da taxa de dissolução, biodisponibilidade e estabilidade destes compostos.
O projeto de processos industriais, sob a ótica da sustentabilidade, visa
alterações substanciais na indústria atual. Para tanto, é exigido o desenvolvimento de novos
processos baseados em matérias-primas renováveis, na utilização mínima necessária de
energia e solventes sem restrições ambientais. Neste contexto, as tecnologias baseadas na
utilização de fluidos pressurizados parecem oferecer soluções para essas demandas.
Tecnologias de extração de antocianinas e carotenóides utilizando fluidos
pressurizados
podem
representar
uma
alternativa
1
ambientalmente
correta
e
economicamente viável em relação aos métodos convencionais de extração onde grandes
quantidades de solventes, longos tempos de extração e altas temperaturas são requeridas, o
que pode promover a degradação destes compostos durante o processo extrativo (JU;
HOWARD, 2003; NOBRE et al., 2006).
Uma vez extraídos os pigmentos, pré-tratamentos destes aditivos podem ser
realizados visando uma maior dissolução e/ou proteção/estabilização destes através de,
respectivamente, técnicas de micronização e encapsulação em matrizes poliméricas
(SELIM; TSIMIDOU; BILIADERIS, 2000). Técnicas utilizando fluidos pressurizados
acima da condição crítica (conhecidos como fluidos supercríticos) permitem os objetivos
requeridos sem levar o aditivo às condições que podem ocasionar sua degradação, isto é,
perda de cor e capacidade antioxidante, condições estas normalmente utilizadas nos
processos convencionais. Adicionalmente, processos de precipitação utilizando fluidos
supercríticos permitem um fácil controle da formação de partículas através de pequenas
variações nas condições de operação (Pressão, Temperatura, Relação Pigmento: Material de
Encapsulação, etc.) (MATTEA; MARTÍN; COCERO, 2008).
Diferentes processos de encapsulação que usam fluidos supercríticos, bem
como equipamentos para a realização destes processos, têm sido desenvolvidos. Estes
processos podem ser classificados de acordo com a função do fluido supercrítico no
processo: solvente [“Rapid Expansion of Supercritical Solutions” (RESS)]; “Supercritical
Solvent Impregnation” (SSI); soluto [“Particles from Gas Saturated Solutions” (PGSS)] ou
anti-solvente [“Supercritical Anti-Solvent” (SAS); “Supercritical Fluid Extraction of
Emulsions” (SFEE)] (MARTÍN; COCERO, 2008).
Geralmente, tanto para os processos de extração com fluidos pressurizados,
quanto para os processos de precipitação, solventes GRAS (“Generally Recognised as
Safe”) são preferidos, sendo dióxido de carbono, água e etanol os mais utilizados. Apesar
de, comercialmente, encontrarem-se disponíveis equipamentos distintos para cada processo,
uma unidade para pesquisa que possibilite o estudo de diferentes processos com fluidos
pressurizados proporcionaria uma melhor relação custo-benefício associada a esta
tecnologia.
2
1.2 Objetivos da Pesquisa
1.2.1 Geral
Construir e validar uma unidade multipropósito para o desenvolvimento de
processos com fluidos pressurizados que possibilite a extração, micronização e
encapsulação de pigmentos funcionais, bem como de outros compostos bioativos em um
único equipamento.
1.2.2 Específicos
o Obter extratos ricos nos pigmentos antocianinas e em outros compostos
antioxidantes a partir de cascas de jabuticaba utilizando a unidade construída para
estudar os seguintes métodos: Extração com Líquidos Pressurizados utilizando
etanol ou água acidificada como solvente; Extração Assistida por CO2 a Alta
Pressão utilizando água acidificada como solvente, comparando os resultados com
os obtidos utilizando o método convencional de Percolação em Leito Fixo;
o Verificar a viabilidade técnica da extração supercrítica assistida por ultrassom na
unidade construída, utilizando a extração de pigmentos carotenóides de sementes de
urucum por CO2 supercrítico como processo de extração modelo;
o Verificar a viabilidade técnica da encapsulação/co-precipitação de extrato rico em
carotenóides oriundo de sementes de urucum, com o polímero polietilenoglicol
(PEG) via SAS, utilizando a unidade construída e CO2 supercrítico como antisolvente;
o Verificar a viabilidade técnica da encapsulação/co-precipitação de extrato
antociânico de casca de jabuticaba e do pigmento funcional também da classe dos
flavonóides rutina, com o polímero polietilenoglicol (PEG) via RESS utilizando a
unidade construída e CO2 supercrítico como solvente e etanol como co-solvente;
o Otimizar o processo de encapsulação/co-precipitação de extrato antociânico de
casca de jabuticaba com o polímero polietilenoglicol (PEG) via RESS, utilizando a
unidade construída e CO2 supercrítico como solvente e etanol como co-solvente,
comparando os resultados relacionados à estabilidade com os obtidos pelo método
3
convencional de gelificação iônica utilizando o biopolímero alginato como material
de encapsulação. Adicionalmente, também se objetivou analisar a influência dos
parâmetros de processo pressão e temperatura na estabilidade do extrato
antociânico.
1.3 Estrutura da Tese de Doutorado
Esta Tese de Doutorado está dividida em 8 capítulos da seguinte forma:
O Capítulo 1 (Introdução) insere o leitor ao tema central desta tese,
colocando, de forma sucinta, os pontos mais relevantes.
O Capítulo 2 (Revisão da literatura) contextualiza o leitor no estado da arte
referente a este trabalho de tese.
O Capítulo 3 detalha a construção e o funcionamento da unidade
multipropósito.
O Capítulo 4 traz o artigo intitulado “Antioxidant pigment extraction using a
home-made pressurized solvent extraction system”, que compara os resultados em termos
de eficiência de extração utilizando CO2 supercrítico puro, obtidos pela unidade
multipropósito com os obtidos por uma unidade de extração supercrítica comercial, com a
finalidade de validação da unidade construída. Para tanto, foram utilizadas como fontes
vegetais modelos duas fontes ricas em pigmentos funcionais: sementes de urucum e cascas
de jabuticaba. Adicionalmente, neste capítulo, a viabilidade técnica de se desenvolver
processos de extrações fracionadas e extrações com fluidos pressurizados assistida com
ultrassom foi verificada. De uma maneira geral, este capítulo relata uma etapa fundamental
para o sucesso no desenvolvimento do restante do trabalho, uma vez que os resultados
obtidos demonstraram que a unidade construída no desenvolvimento de extrações
utilizando fluidos pressurizados produz resultados similares aos obtidos por equipamentos
comerciais, bem como resultados reprodutíveis.
O trabalho seguinte foi explorar a potencialidade de obtenção de extratos ricos
em antocianinas utilizando a unidade multipropósito validada conforme descrito no capítulo
anterior. Neste contexto, o artigo apresentado no Capítulo 5, intitulado “Pressurized liquid
4
extraction of phenolic compounds from jabuticaba skins: optimization study”, investiga a
influência das variáveis: Pressão, Temperatura e Tempo de Extração Estática, e otimiza o
processo de extração de antocianinas e outros compostos fenólicos de cascas de jabuticaba
utilizando etanol pressurizado. Ao final, os resultados obtidos empregando condições
otimizadas foram comparados aos obtidos utilizando o método convencional à baixa
pressão de percolação em leito fixo.
O artigo apresentado no Capítulo 6, intitulado “Optimization of bioactive
compounds extraction from jabuticaba (Myrciaria cauliflora) skins assisted by high
pressure CO2”, traz os resultados obtidos para a extração de antocianinas e outros
compostos fenólicos de cascas de jabuticaba também utilizando a unidade construída,
porém agora configurada para desenvolver uma metodologia de extração diferente,
chamada de extração assistida com CO2 a alta pressão. Neste trabalho é investigada a
influência das variáveis: Pressão, Temperatura e Relação entre o Volume de Matéria-prima
+ Solvente e o Volume de CO2 dentro da célula extratora na extração de antocianinas e
outros compostos fenólicos. Ao final, foi realizada uma comparação dos resultados obtidos
empregando condições otimizadas com os obtidos desenvolvendo a metodologia de
extração com líquidos pressurizados também utilizando esta unidade com o mesmo
solvente (Água acidificada) nas mesmas temperatura e pressão.
Continuando com a mesma abordagem descrita no Capítulo 4, de verificar se a
unidade construída produz resultados similares aos obtidos por equipamentos similares
(validação da unidade multipropósito construída), o Capítulo 7 traz o artigo intitulado
“Micronization and encapsulation of functional pigments using supercritical carbon
dioxide” que apresenta um estudo comparativo das partículas formadas pela unidade
construída com as formadas por outros equipamentos de outros grupos de pesquisa.
Adicionalmente, neste capítulo foi estudada a viabilidade técnica de se desenvolver
processos de formação de partículas encapsuladas de extrato de sementes de urucum
(obtidos conforme descrito no Capítulo 4), de extrato de cascas de jabuticaba (obtidos
conforme descrito no Capítulo 5) e de pigmento rutina puro utilizando como material de
encapsulação polietilenoglicol (PEG).
5
Uma vez demonstrada a potencialidade de formação de partículas encapsuladas
de extrato de cascas de jabuticaba empregando a unidade multipropósito no Capítulo 7 este
processo foi mais bem explorado no Capítulo 8, bem como foi verificado o potencial de
aumento de estabilidade deste extrato quando protegido por matrizes poliméricas. O artigo
intitulado “Stabilization of anthocyanins from jabuticaba skins by encapsulation using
supercritical CO2” analisa também a influência dos parâmetros de processo pressão e
temperatura na estabilidade do extrato antociânico. Ao final, foi realizada uma comparação
dos resultados obtidos empregando condições otimizadas com os obtidos utilizando o
método convencional à baixa pressão gelificação iônica. De uma maneira geral, este
capítulo encerra os objetivos previstos para este trabalho de tese, fechando o ciclo de
validação-experimentação dos processos de extração e formação de partículas que a
unidade multipropósito pode desenvolver.
O Capítulo 9 (Conclusões gerais) discorre sobre os principais resultados
obtidos em cada um dos artigos apresentados nesta tese.
O Apêndice I traz um artigo de revisão sobre as antocianinas, discorrendo
sobre a estrutura química, propriedades benéficas à saúde e principais fontes de obtenção
destes compostos, enfatizando que para o Brasil as cascas de jabuticaba podem ser uma
potencial fonte destes pigmentos instáveis. Já o Apêndice II traz o detalhamento do
procedimento de cálculo para a determinação da concentração de antocianinas e compostos
fenólicos presentes nos extratos obtidos neste trabalho. O Apêndice III traz um artigo de
revisão sobre os métodos de formação de partículas que discorre sobre as vantagens e
limitações dos principais métodos empregados atualmente, descrevendo como cada
processo é desenvolvido, bem como enfatizando a potencialidade de utilizar os métodos
que empregam fluidos supercríticos na formação de partículas encapsuladas em matrizes
poliméricas. Finalmente, o Apêndice IV traz o manual de operação da unidade
multipropósito construída, detalhando o procedimento adotado para o desenvolvimento dos
processos estudados nesta tese, bem como os procedimentos para a realização de outros
processos que a unidade possibilita desenvolver.
6
Nesta Tese de Doutorado, o desenvolvimento dos capítulos e apêndices
apresenta-se através de artigos publicados/submetidos ou a serem submetidos a periódicos.
Permissões para a utilização dos artigos publicados em periódicos foram devidamente
obtidas com as respectivas editoras, previamente.
7
8
CAPÍTULO 2 - REVISÃO BIBLIOGRÁFICA
2.1. Pigmentos Funcionais
Existe grande interesse sobre as propriedades benéficas que os componentes
nutracêuticos presentes em alimentos propiciam à saúde humana (PALIYATH, 2003). Têm
sido adquirido conhecimento científico a respeito dos ingredientes presentes nos alimentos
que ingerimos, com potencial para prevenir e tratar doenças específicas. Paralelo a isto,
novas tecnologias, como a biotecnologia e a engenharia genética especificamente, têm
criado uma era onde descobertas científicas e produtos inovadores são cada vez mais
comuns. Estes desenvolvimentos têm resultado num aumento do número de produtos
potencialmente nutricionais com benefícios para a saúde, produtos estes chamados de
“alimentos funcionais”. Os alimentos funcionais, além do valor nutritivo básico, contêm um
equilíbrio próprio de ingredientes, os quais podem ajudar diretamente na prevenção e
tratamento de doenças (GOLDBERG, 1994). As substâncias bioativas presentes nos
alimentos funcionais representam constituintes “extranutricionais” naturalmente presentes
em pequenas quantidades na matriz do alimento (KITTS, 1994).
Estas substâncias bioativas são pertencentes ao grupo dos compostos do
metabolismo secundário de plantas, também chamados de compostos fitoquímicos, e
podem ser definidos como substâncias derivadas de plantas que são altamente ativas do
ponto de vista nutricional, fisiológico e/ou medicinal (GOLDBERG, 1994).
Os pigmentos naturais presentes naturalmente em alimentos proporcionam cor,
contribuindo para seu aspecto visual, atributo que está diretamente relacionado à aceitação
deste alimento pelos consumidores (CLYDESDALE, 1993). O consumo de um alimento
que possui em sua composição aditivos corantes naturais é associado à imagem de um
alimento de qualidade e saudável; além dos corantes sintéticos possuírem a tendência de
serem vistos como indesejáveis e prejudiciais, alguns são responsabilizados por reações
alergênicas e de intolerância (MONTES et al., 2005). Pigmentos das classes dos
flavonóides e carotenóides têm sido relacionados a importantes funções e ações
fisiológicas, podendo ser considerados promotores da saúde humana. A ingestão de frutas e
vegetais está sendo associada com a diminuição do desenvolvimento de doenças crônico9
degenerativas tais como câncer, inflamações, doenças cardiovasculares, catarata,
degeneração macular, entre outras (KONG et al., 2003; KRINSKY, 1994). De forma geral,
as propriedades antioxidantes destes compostos parecem ser a chave para a elucidação dos
mecanismos envolvidos nestas ações.
2.1.1 Flavonóides
Os flavonóides são pigmentos naturais presentes nos vegetais que desempenham um
papel fundamental na proteção contra agentes oxidantes, como por exemplo, os raios
ultravioleta, a poluição ambiental, substâncias químicas presentes nos alimentos, entre
outros. Eles atuam como agentes terapêuticos num elevado número de patologias, tais como
arteriosclerose, cancer, etc (PASSAMONTI et al., 2009).
Dado que não podem ser sintetizados pelo nosso organismo, sendo
representativos da parte não energética da dieta humana, os flavonóides são obtidos através
da ingestão de alimentos que os contenham. Exemplos de fontes de flavonóides são frutas,
verduras, cerveja, vinho, chá e soja. A maioria dos flavonóides presentes no vinho provém
da uva, especialmente da pele (KOSMIDER; OSIECKA, 2004).
Os flavonóides apresentam uma estrutura química base C6-C3-C6 (dois anéis
benzênicos – A e B – ligados através de um anel pirano – C (Figura 2.1.1.1). Dependendo
da substituição e do nível de oxidação no anel C, os flavonóides podem ser divididos em 14
classes, sendo os que se incluem na dieta humana divididos essencialmente em 6 grupos
(PASSAMONTI et al., 2009; KOSMIDER; OSIECKA, 2004; ERLUND, 2004):
• Flavanóis – possuem um grupo hidroxila na posição 3. Exemplos: catequina,
epicatequina.
• Flavonóis – possuem um grupo carbonila na posição 4, um grupo hidroxila
na posição 3 e uma ligação dupla entre as posições 2 e 3. Exemplos:
quercetina, kaempferol, quercitagetina, etc.
• Flavonas – possuem um grupo carbonila na posição 4 e uma ligação dupla
entre as posições 2 e 3. Exemplos: rutina, apigenina, luteoleína, etc.
10
• Antocianidinas – possuem um grupo hidroxila na posição 3 e duas ligações
duplas: uma entre o átomo de oxigênio e o carbono 2 e outra entre os
carbonos 2 e 3. Exemplos: cianidinas, petunidinas, malvidina, etc.
• Isoflavonóides – possuem um grupo carbonila na posição 4 e o anel B
encontra-se ligado ao restante da molécula através do carbono 3. Podem
ainda possuir uma ligação dupla entre os carbonos 2 e 3. Exemplos:
genisteína, coumestrol, etc.
• Flavononas – possuem um grupo carbonila na posição 4. Exemplos:
miricetina, hesperidina, etc.
Figura 2.1.1.1 - Estrutura base dos flavonóides.
Dentro da mesma classe, os flavonóides diferem na substituição dos anéis A e
B. Estes se encontram na natureza sob a forma de glicosídeos, o que promove uma melhor
absorção intestinal e uma maior biodisponibilidade destes compostos. Os glicosídeos
formam-se através da união de resíduos de D-glicose à posição 3 ou à posição 7 destes
flavonóides, sendo a primeira substituição a mais freqüente. Outros resíduos de açúcares,
que também podem se encontrar ligados a este tipo de compostos, são a D-galactose, a Lramnose, a L-arabinose, a D-xilose e o ácido D-glucurônico (ERLUND, 2004).
As antocianinas são um dos exemplos de flavonóides encontrados na natureza
sob a forma de glicosídeos. Antocianinas são compostos derivados das antocianidinas. Nas
antocianinas, uma ou mais hidroxilas das posições 3, 5 e 7 estão ligadas a açúcares, aos
11
quais podem estar ligados ácidos fenólicos. Os diferentes grupos R e R´ ligados nas
posições 3´e 5´ e açúcares ligados nas posições 3, 5 e 7, assim como os ácidos a eles
ligados, caracterizam os diferentes tipos de antocianinas, sendo que a mais comum é a
cianidina-3-glicosídeo (SANTOS; MEIRELES, 2009). Maiores detalhes sobre as
antocianinas: estrutura química, instabilidade, propriedades benéficas à saúde e principais
fontes de obtenção destes compostos são encontrados no Apêndice I desta tese. Neste
Apêndice também é demonstrado que, para o Brasil, as cascas de jabuticaba (Figura
2.1.1.2) podem ser uma potencial fonte destes pigmentos.
Diversos fatores, tais como luz, temperatura, pH, entre outros, desencadeiam a
degradação oxidativa das antocianinas, sendo estes compostos mais estáveis em meios
ácidos do que em alcalinos. A natureza da estrutura iônica das antocianinas possibilita
mudanças na estrutura molecular de acordo com o pH, resultando diferentes cores. Em pH
< 3, a antocianina cianidina-3-glicosídeo, por exemplo, existe primariamente como uma
estrutura molecular que resulta na cor vermelha. Aumentando-se os valores de pH do meio,
novas formas moleculares são produzidas, resultando em cores que vão desde o violeta a
uma forma incolor. Zhang et al. (2010) sugere que a degradação da cianidina-3-glicosídeo
se inicia devido a facilidade de se hidrolisar o grupo glicosídeo ligado a estrutura base. Um
esquema das transformações estruturais das antocianinas em função do pH gerando
soluções coloridas é apresentado no Apêndice II.
Figura 2.1.1.2 - Cascas de jabuticaba.
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Além da identificação dos tipos de antocianinas presentes nos extratos, a
determinação da sua concentração, bem como de outros compostos co-extraídos, é de
extrema importância, pois elas estão associadas às funções biológicas destes extratos. Para
a determinação da concentração dos flavonóides antocianinas nos extratos, os métodos
baseados nas transformações estruturais das antocianinas em função do pH, gerando
soluções coloridas, têm propiciado resultados confiáveis, sendo o método denominado de
método do pH diferencial descrito por Giusti e Wrolstad (2001) um dos mais utilizados.
Maiores informações sobre o procedimento de cálculo deste método são encontradas no
Apêndice II desta tese. Neste Apêndice também é descrito como é feito o cálculo da
concentração de compostos fenólicos presentes nos extratos (incluindo a concentração de
antocianinas e outros compostos fenólicos co-extraídos).
2.1.2 Carotenóides
Os carotenóides são uma classe de compostos lipofílicos amplamente
conhecidos pelo seu poder corante que pode variar do amarelo ao vermelho. A sua estrutura
básica é um tetraterpeno com 40 átomos de carbono, formado por oito unidades
isoprenóides de cinco carbonos, ligados de tal forma que a molécula é linear com simetria
invertida no centro. A principal característica dos carotenóides é um sistema de ligações
duplas conjugadas, que corresponde ao cromóforo, e que permite a estes compostos
absorver luz na região do visível, como pode ser observado na estrutura do β-caroteno
(Figura 2.1.2.1) (MIKI, 1991).
Figura 2.1.2.1 - Estrutura do β-caroteno
13
Os carotenóides são inicialmente divididos em dois grandes grupos, os
carotenos que quimicamente são hidrocarbonetos, e as xantofilas que são derivados
oxigenados. Neste último grupo estão incluídos pigmentos que possuem em sua estrutura
grupos hidroxílicos, carbonílicos, carboxílicos e/ou epóxidos. Podendo ser também
acíclicos, monocíclicos ou bicíclicos. Muitas outras modificações estruturais ainda são
possíveis, permitindo obtenção de uma diversidade de compostos (BRITTON, 1995).
Alguns dos tipos de carotenóides mais amplamente distribuídos na natureza são: βcaroteno, licopeno, luteína, zeaxantina, bixina, entre outros (MCCLEMENTS et al., 2009;
ALBUQUERQUE; MEIRELES, 2010).
A significância dos carotenóides não é somente devido às suas propriedades
corantes, eles também são muito importantes para a saúde. Estes compostos também
desempenham importante papel nutricional como precursores de vitamina A, além de
outras ações, tais como proteção contra alguns tipos de câncer, doenças cardiovasculares,
cataratas, degeneração macular e melhoria do sistema imunológico (GOUVEIA; EMPIS,
2003; ROBERT et al., 2003).
Edge, McGarvey e Truscott (1997) associaram, em seus estudos, as funções
biológicas dos carotenóides em seres humanos. De acordo com estes autores, esses
compostos podem ser importantes na proteção de células atuando como antioxidantes
contra radicais livres e seqüestrando oxigênio singlete devido ao seu longo sistema de
ligações duplas conjugadas.
Pigmentos das classes dos carotenóides, assim como os da classe dos
flavonóides, têm sua degradação oxidativa acelerada pela luz, temperatura e/ou pH extremo
na presença de oxigênio (SANTOS; MEIRELES, 2010).
Comercialmente, uma das fontes vegetais de pigmentos mais amplamente
utilizadas pela indústria de alimentos é o urucum (Bixa orellana), correspondendo em torno
de 90% do total de consumo de corantes naturais no Brasil, e em torno de 70% de corantes
naturais mundialmente empregados em alimentos (CONSTANT; STRINGHETA; SANDI,
2002). No Brasil, o urucum é uma espécie economicamente importante e ocorre em todas
as regiões brasileiras e sua disseminação em diversas regiões do mundo está relacionada
14
com a procura por corante natural pelas indústrias de medicamentos, cosméticos, têxteis e
principalmente alimentar (FRANCO et al., 2008).
Urucum (Bixa orellana L.) é uma arvore tropical cujas sementes são ricas no
pigmento da classe dos carotenóides conhecido como bixina, o qual é somente encontrado
nesta fonte vegetal. Quimicamente bixina (C25H30O4) é um mono-metil ester do ácido
dicarboxílico norabixina, um outro importante pigmento encontrado nas sementes de
urucum (Figura 2.1.2.2). Entre outras aplicações, estes corantes são usados pela indústria
alimentícia para melhorar queijos, margarinas e manteigas (ANDERSON et al., 1997).
Figura 2.1.2.2 - Sementes de urucum.
2.2. Métodos de Extração
Existem, na literatura, várias metodologias de obtenção de extratos de vegetais,
utilizando vários solventes, empregando ou não altas pressões e temperaturas ou até mesmo
associando metodologias no intuito de obter um ou mais extratos com o perfil fitoquímico
desejável, com alto rendimento e que não possua características indesejáveis como a
presença de compostos co-extraídos como ceras e clorofila e a presença residual de
solventes (BRAGA, 2005). A seguir são descritos alguns métodos que foram utilizados na
obtenção de extratos ricos em antocianinas e no carotenóide bixina.
O solvente mais utilizado na extração de antocianinas é o metanol, apesar da
sua toxicidade. Entretanto, para algumas aplicações onde o aspecto quantitativo não é
15
prioritário, etanol e água também têm sido utilizados. A limitação do uso de etanol e água
está relacionada com a menor eficiência na extração de antocianinas (TERCI, 2004). Estas
metodologias, também chamadas de convencionais, implicam na co-extração de compostos
não fenólicos como açúcares, ácidos orgânicos e proteínas, requerendo uma subseqüente
purificação destes extratos, uma vez que estes processos geralmente apresentam baixa
seletividade (CASTAÑEDA-OVANDO et al., 2009).
Comumente, métodos de extração convencionais, tais como soxhlet, por
agitação, por percolação em leito fixo, entre outros, são utilizados nos processos de
extração de antocianinas, entretanto, estes métodos geralmente consomem muito tempo e
grande quantidade de solventes e podem degradar estes compostos durante o processo
extrativo. Segundo Pinedo (2007), a hidrólise completa dos açúcares ligados a antocianinas,
açúcares que propiciam melhor estabilidade dos pigmentos após a extração, ocorre em 1
hora a 333,15 K na presença de etanol acidificado com 1% de HCl.
Visando eliminar estas limitações, métodos de extrações rápidas que utilizam
fluidos pressurizados têm sido empregados com sucesso para a obtenção de extratos ricos
em antocianinas (ARAPITSAS; TURNER, 2008). Também, a utilização de tratamentos
com ultrassom e CO2 a alta pressão, auxiliares ao processo extrativo de antocianinas, tem
demonstrado melhorar o desempenho da extração, aumentando rendimento, reduzindo o
tempo de extração, etc. (CAI et al., 2003; XU et al., 2010).
Extratos comerciais de sementes de urucum são obtidos utilizando vários
processos, tais como extração com óleos vegetais, com solução alcalina e com solventes
orgânicos: clorofórmio, diclorometano, acetona e metanol (PRESTON; RICKARD, 1980).
As preparações de extratos ricos em pigmentos têm as desvantagens de conterem pequenas
concentrações do composto bioativo alvo (bixina), bem como apresentar solvente tóxico no
produto final. Suspensões de extratos de urucum em óleos vegetais são mais concentradas,
mas podem conter produtos de degradação devido ao fato de altas temperaturas (> 100 ºC)
serem empregadas nestes processos, afetando a qualidade do extrato (MCKEOWN;
MARK, 1962; REITH; GIELEN, 1971).
16
Visando eliminar estas limitações, o emprego de fluidos pressurizados em
condições supercríticas como solventes de extração na obtenção de extratos ricos no
carotenóide bixina tem se demonstrado uma potencial opção (MENDES et al., 2003).
A alta seletividade do processo de extração supercrítica, através de pequenas
alterações nas variáveis do processo (pressão e temperatura), aliada à eliminação de
solventes residuais no produto final e ao uso de baixas temperaturas ao se utilizar CO2
supercrítico puro na obtenção de extratos ricos em bixina a partir de sementes de urucum,
têm sido as principais justificativas para a utilização deste método de extração em
substituição aos métodos convencionais (ANDERSON et al., 1997; NOBRE et al., 2006;
SILVA et al., 2008).
2.2.1 Extração com Fluidos Pressurizados
Os fluidos supercríticos caracterizam-se por a sua temperatura e pressão serem
superiores aos correspondentes valores críticos. Acima do ponto crítico, deixa de haver
tensão superficial e separação entre as fases líquida e gasosa em equilíbrio, formando-se
uma única fase supercrítica, cujas propriedades são intermediárias daqueles dois estados.
Abaixo do ponto crítico o fluido pode existir como um líquido ou como um vapor
(SANDLER, 1989). A Tabela 2.2.1.1 registra a pressão e temperatura críticas de alguns
fluidos com interesse em processos de extração.
Tabela 2.2.1.1 - Propiedades críticas de alguns fluidos com interesse em extração (MENDES
et al., 2003)
Fluido
Tc (ºC)
Pc (bar)
50,4
Etileno
9,4
Dióxido de carbono
31,1
Etano
32,4
Óxido nitroso
36,6
Propano
96,8
17
73,8
48,8
72,4
42,5
Etanol
240,9
Benzeno
289,1
Tolueno
318,8
Água
401,3
61,4
48,9
41,0
221,2
Nas proximidades da região crítica, os fluidos não apenas são solventes eficazes
para a extração de compostos bioativos de fontes vegetais, mas apresentam uma série de
peculiaridades que os tornam mais vantajosos com relação aos solventes líquidos,
comumente utilizados (REZENDE, 1998). Algumas peculiaridades/vantagens são:
* Ausência de resíduos do solvente nos produtos (extratos);
* Uma variedade maior de solventes pode ser utilizada, já que as características
básicas da extração supercrítica devem-se, além das propriedades do solvente, às condições
termodinâmicas;
* A seletividade de um dado soluto, em uma solução do solvente, pode ser
controlada, manipulando-se a densidade do solvente.
2.2.1.1 Extração com Líquidos Pressurizados
Um fluido pressurizado e aquecido à pressões e temperaturas abaixo das críticas
pode existir como um líquido ou como um vapor. No caso de o fluido ser etanol ou água,
por exemplo, à pressões menores que 200 bar e temperaturas menores que 200 ºC este
fluido se mantém no estado líquido, apesar de a temperatura estar acima da sua temperatura
de ebulição a pressão atmosférica. A utilização destes fluidos nestas condições em
processos extrativos permite uma melhora da solubilidade dos compostos a serem
extraídos, bem como uma aceleração da cinética de dessorção destes compostos da matriz
vegetal (RICHTER et al., 1997).
Nestes processos, a utilização de elevada temperatura de extração aumenta a
capacidade do solvente de solubilizar os compostos alvo, já o emprego de altas pressões
18
acelera a difusão nos poros da matriz e diminui a viscosidade do solvente, causando uma
maior penetração do solvente na matriz e, portanto aumentando sua capacidade de extração
(LOPEZ-AVILA, 1999).
O método de extração com líquidos pressurizados emprega menor volume de
solvente que as extrações convencionais (soxhlet, etc.) e é mais rápido (tempo de extração
varia entre 10 e 20 minutos).
De outras perspectivas, incluindo considerações sobre meio ambiente,
segurança e economia, muitos esforços têm sido feitos com o objetivo de diminuir o uso de
solventes orgânicos nos laboratórios químicos e processos industriais. O uso da água como
solvente de extração apresenta vantagens adicionais. A água é barata, não-tóxica, não
combustível ou explosiva e é ambientalmente segura. A alta temperatura, o aumento do
produto iônico e a mudança na constante dielétrica fazem com que os compostos bioativos
a serem extraídos sejam altamente solúveis na água sob estas condições. A constante
dielétrica pode influenciar as taxas de extrações com o aumento da polaridade do solvente
extrativo. Assim, a densidade pode ser usada para manipular a constante dielétrica e,
conseqüentemente as cinéticas de extração (CORRALES; BUTZ; TAUSCHER, 2008; JU;
HOWARD, 2003; CACACE; MAZZA, 2002).
Mais detalhes sobre a metodologia de extração com etanol e água
pressurizados, particularmente de pigmentos antociânicos, são apresentados nos Capítulos 5
e 6, respectivamente.
2.2.1.2 Extração com CO2 supercrítico
Na maioria das vezes tem-se utilizado o dióxido de carbono para a extração
supercrítica de produtos naturais. A grande aceitação do dióxido de carbono deve-se
(REVERCHON; OSSÉO, 1994):
* À sua atoxidade, em pequenas quantidades;
* À sua não-inflamabilidade;
* Ao seu ponto crítico ocorrer em condições relativamente brandas. A
temperatura crítica é de 31,0 °C (304,1 K) e a pressão crítica de 73,8 bar (Figura 2.2.1.2.1);
19
* À sua estabilidade química;
* À sua disponibilidade a baixo custo. O dióxido de carbono pode ser obtido,
por exemplo, a partir de processos fermentativos.
Mais detalhes sobre a metodologia de extração com CO2 supercrítico,
particularmente de pigmento bixina da classe dos carotenóides, são apresentados no
Capítulo 4.
Figura 2.2.1.2.1 - Diagrama de fase pressão-temperatura do dióxido de carbono.
2.2.2 Métodos de Extração Assistida
2.2.2.1. Com ultrassom
Processos de extração assistidos com ultrassom têm sido cada vez mais
estudados, pois o emprego do ultrassom na extração de diferentes fontes vegetais utilizando
diferentes solventes de extração tem propiciado: i) um maior rendimento de extração; ii)
um aumento da taxa de extração; iii) uma redução no tempo de extração, dentre outras
vantagens (VILKHU et al., 2008).
A gama de aplicações do uso da extração assistida com ultrassom inclui ervas,
óleos, proteínas e compostos bioativos de materiais vegetais e animais (por exemplo,
20
compostos fenólicos, antocianinas, compostos aromáticos, polissacarídeos, etc.) e a gama
de solventes de extração já empregados nestes processos incluem água, etanol, hexano,
dióxido de carbono supercrítico, entre outros (VINATORU, 2001; RIERA et al., 2010;
ROMDHANE; GOURDAN, 2002).
A melhora do processo de extração através do uso do ultrassom tem sido
atribuída à propagação das ondas de pressão ultrasônicas, o que resulta em cavitação. Altas
forças de cisalhamento provocam o aumento da transferência de massa dos compostos a
serem extraídos. Durante esta etapa temperaturas muito elevadas (cerca de 5.000 K) e
pressões (aproximadamente 2.000 bar) são atingidas localmente. Quando as bolhas atingem
um volume em que já não podem absorver energia, ocorre um colapso violento. Este
fenômeno é denominado de cavitação. A implosão das bolhas de cavitação gera
microturbulência, colisões entre as partículas em alta velocidade e perturbação nos microporos das partículas da matriz vegetal, o que acelera a difusão. Este efeito proporciona uma
exposição de novas superfícies aumentando ainda mais a transferência de massa (JIANBING et al., 2006).
Os efeitos do ultrassom no processo de extração dependem da freqüência e
capacidade do equipamento e do tempo empregado para a extração. Alguns autores têm
informado a ocorrência de transformações químicas nos extratos, resultantes da utilização
de longos tempos de irradiação ultrasônica (VINATORU, 2001).
Mais detalhes sobre a metodologia de extração assistida com ultrassom são
apresentados no Capítulo 4.
2.2.2.2. Com CO2 a alta pressão
O efeito explosivo da descompressão de CO2 a alta pressão em primeiro lugar
demonstrou romper células bacterianas através da rápida liberação de pressão de gás com o
objetivo de recolher o conteúdo da célula (lise celular). Numerosos estudos têm mostrado a
eficácia do uso de CO2 a alta pressão para inativar microrganismos e enzimas (BALABAN
et al., 1991; KINCAL et al., 2006).
21
Devido à similaridade entre os processos que visam à inativação microbiana e
os que visam à extração sólido-líquido de compostos bioativos de matrizes vegetais, Xu et
al. (2010), pela primeira vez, deduziram e confirmaram com seus resultados experimentais
que extrações sólido-líquido, assistidas com CO2 a alta pressão, poderiam propiciar
melhores resultados que quando não assistidas.
A melhora do processo de extração através do uso de CO2 a alta pressão foi
atribuída às habilidades do CO2 a alta pressão de modificar a membrana celular, diminuir o
pH intracelular, desorientar o equilíbrio eletrolítico intracelular, remover os componentes
vitais das células e membranas celulares (XU et al., 2010). Adicionalmente, no caso
especificamente estudado por Xu et al. (2010), o uso de CO2 a alta pressão melhorou o
processo de extração possivelmente por este ter reagido com o solvente utilizado (água),
gerando um produto de reação (água carbonatada) com mais habilidades em extrair os
compostos bioativos desejados.
Mais detalhes sobre a metodologia de extração assistida com CO2 a alta
pressão, particularmente de pigmentos antociânicos, são apresentados no Capítulo 6.
2.3. Formação de Partículas
A aplicação dos pigmentos das classes dos flavonóides e carotenóides como
aditivos em produtos alimentícios é severamente limitada: por sua rápida degradação
acelerada pela luz, temperatura e presença de oxigênio, por sua baixa solubilidade em
sistemas aquosos, etc. (ÖZEN; AKBULUT; ARTIK, 2009; MATTEA; MARTÍN;
COCERO, 2009). A redução do tamanho das partículas destes pigmentos funcionais e/ou
sua encapsulação em geral tem sido opções que visam eliminar estas limitações (SUO et al.,
2005; MATTEA; MARTÍN; COCERO, 2009).
A produção controlada de partículas de um determinado produto (que pode ser
um aditivo alimentício como corante) cujas características físicas (tamanho, morfologia e
estrutura) estão otimizadas é designada por “engenharia de partículas”. O objetivo principal
desta disciplina é a incorporação de atributos desejáveis nas partículas, tais como uma
22
estreita distribuição de tamanhos, elevada estabilidade, maior biodisponibilidade,
distribuição controlada e administração dirigida (CHOW et al., 2007).
Diferentes métodos de formação de partículas nas formas puras ou
encapsuladas têm sido desenvolvidos. Maiores detalhes sobre estes métodos: vantagens e
limitações são apresentadas no Apêndice III desta tese. Dentre os novos métodos de
formação de partículas, particular ênfase se é dada aos processos RESS [Rapid Expansion
of Supercritical Solutions] e SAS [Supercritical Anti-Solvent] que utilizam CO2
supercrítico na função de solvente e anti-solvente, respectivamente.
Além do controle das propriedades das partículas, estes novos processos trazem
vantagens competitivas aos métodos tradicionais, pois requerem menor manuseamento, o
que permite um aumento do rendimento e simplifica os procedimentos de limpeza e
esterilização, e estão mais aptos para o aumento de escala. Estes processos podem também
operar em modo contínuo, ao contrário da grande maioria dos processos convencionais, que
operam em descontínuo (SANTOS; MEIRELES, 2010). Mais detalhes sobre cada um
deles, bem como a potencialidade de utilizar estes métodos que empregam fluidos
supercríticos na formação de partículas encapsuladas em matrizes poliméricas visando à
estabilização de pigmentos funcionais, particularmente os da classe dos carotenóides,
também é encontrado no Apêndice III.
Mais detalhes sobre a metodologia de formação de partículas nas formas puras
ou encapsuladas empregando os processos RESS [Rapid Expansion of Supercritical
Solutions] e SAS [Supercritical Anti-Solvent] são apresentados no Capítulo 7. Detalhes
sobre a estabilização destes pigmentos funcionais, particularmente os antociânicos, por
encapsulação em matrizes poliméricas, são encontrados no Capítulo 8.
2.4. Equipamentos que utilizam fluidos pressurizados
2.4.1 Para Extração
2.4.1.1 Para Extração com Líquidos Pressurizados
23
A extração com líquidos pressurizados, também conhecida como extração com
solventes pressurizados, extração acelerada com solventes ou extração com fluidos
pressurizados é desenvolvida em equipamentos comerciais e “Home-made”.
Existem diferentes fornecedores de sistemas de extração para extração com
líquidos pressurizados. Dionex oferece dois sistemas de extração: 1) ASE® 150 - um
sistema de extração com uma célula de extração simples (Figura 2.4.1.1.1a); 2) ASE® 350 um sistema de extração que possibilita a extração automatizada de até 24 amostras (Figura
2.4.1.1.1b). Fluid Management Systems oferece um único sistema de extração seqüencial
capaz de extração automatizada de até 6 amostras (Figura 2.4.1.1.2).
Figura 2.4.1.1.1 - Equipamento comercial para extração com líquidos pressurizados. a)
ASE® 150; b) ASE® 350 (DIONEX, 2010)
Figura 2.4.1.1.2 - Equipamento comercial para extração com líquidos pressurizados
(FLUID MANAGEMENT SYSTEMS, 2010)
24
2.4.1.2 Para Extração com CO2 Supercrítico
Em 1978 começou a operar o primeiro processo industrial de extração de
cafeína com CO2 supercrítico. A extração supercrítica já é utilizada em escala industrial
para descafeinação do café e de chá e também para a extração de lúpulo (BRUNNER,
1994).
Estima-se que há atualmente cerca de 250 plantas industriais que utilizam esta
1
tecnologia . Segundo Perrut (2000), existem em operação unidades de extração
supercríticas na Itália e na França que operam no processamento de ingredientes de
alimentos, princípios ativos farmacêuticos e cosméticos. Também há relatos que
recentemente na Espanha, o primeiro processo de extração com CO2 supercrítico começou
a operar em San Vicente de Alcántara (Badajoz) em 2005 e em 2008, a empresa começou a
operar outra planta em Valência1. Entretanto, não existe nenhuma unidade de produção em
escala industrial na América do Sul, exceto por uma unidade semi-piloto que se encontra
em testes2.
Existem diferentes fornecedores que oferecem equipamentos para o
desenvolvimento de extrações com fluidos supercríticos com diferentes configurações e
capacidades, porém a utilização de equipamentos “Home-made” também é muito usual em
nível laboratorial. A série Spe-ed SFE (Supercritical Fluid Extraction) da Applied
Separations é uma das séries mais populares, pois oferece uma variedade de opções que
atendem necessidades analíticas e de pesquisa. O sistema de aquecimento da célula
extratora constitui de um forno, o que permite que células extratoras de diferentes
capacidades possam ser utilizadas nos processos de extração, ou mais de uma célula
extratora seja utilizada, o que possibilita processos de extração de modo semi-contínuo
(enquanto uma célula extratora é empacotada a outra está sendo utilizada e vice-versa)
(Figura 2.4.1.2.1). Adicionalmente, ao equipamento Spe-ed SFE uma bomba de co-solvente
1
2
Informação pessoal de Prof. Dra. M. J. Cocero.
Informação pessoal de A. Astini.
25
pode ser incorporada (módulo comprado a parte) para possibilitar a extração com CO2
supercrítico + co-solventes.
Figura 2.4.1.2.1 - Equipamento comercial para extração com CO2 supercrítico (APPLIED
SEPARATIONS, 2010)
2.4.1.3 Para Extração assistida com ultrassom
Recentemente, o design de equipamentos com transdutores de ultrassom tem
avançado para fornecer capacidade de processamento em nível industrial. Atualmente, 16
kW é a potência do maior transdutor de ultrassom disponível. Fornecedores industriais de
transdutores de ultrassom têm promovido cada vez mais a utilização de ondas ultrasônicas
em diversos processos de extração convencionais para diferentes aplicações nos últimos
anos (VILKHU et al., 2008).
Vários projetos de extratores com transdutores de ultrassom acoplados têm sido
descritos por Chisti (2003) e Vinatoru (2001). Estes incluíram transdutores de ultrassom ora
26
no agitador mecânico que fica em contato com meio extrativo (solvente + material a ser
extraído) (Figura 2.4.1.3.1) ora na parede das células de extração.
Figura 2.4.1.3.1 - Equipamento comercial para extração assistida com ultrassom
(HIELSCHER, 2010)
Sistemas modernos de ultrassom possuem um dispositivo automatizado que
ajusta a freqüência das ondas ultrasônicas para assegurar que a potência máxima é
transmitida ao conteúdo da célula extratora, bem como possibilitam exposição às ondas
ultrasônicas de modo intermitente ou contínuo (ROMDHANE; GOURDAN, 2002).
2.4.1.4 Para Extração assistida com CO2 a alta pressão
Há somente um relato do processo de extração assistida com CO2 a alta pressão,
tendo este sido desenvolvido em equipamento “Home-made” projetado para preservar
alimentos por inativação microbiana (Xu et al., 2010). Segundo Parton et al. (2007) ainda
não há equipamentos comerciais disponíveis de pasteurização/inativação bacteriana de
alimentos que utiliza CO2 a alta pressão devido, principalmente, a duas razões: i) falta de
conhecimento sobre instalações de alta pressão para processamento de alimentos; ii) pobre
27
compreensão do mecanismo de inativação induzido pela pressão do CO2 sobre os
microrganismos.
2.4.2 Para Formação de Partículas
2.4.2.1 Para Formação de Partículas Via SAS
Processos de formação de partículas via SAS são desenvolvidos tanto em
equipamentos comerciais como em “Home-made”. A empresa Thar Technologies,
acompanhando o desenvolvimento científico produzido por pesquisadores que estudam
processos de formação de partículas via SAS, lançou recentemente no mercado um produto
que pode realizar o processo SEDS (Solution Enhanced Dispersion by Supercritical fluids),
um subtipo do processo SAS (Figura 2.4.2.1.1). Sabe-se que anteriormente a esta
modificação os pesquisadores, que adquiriram os equipamentos para o desenvolvimento
dos processos via SAS convencional, já faziam a modificação em seus laboratórios
(JACOBSON et al., 2010). Basicamente a modificação se dá na entrada da tubulação de
CO2 através da inserção de uma conexão “T” que permite a entrada da solução contendo a
substância ativa a ser micronizada ou encapsulada coaxialmente ao CO2 supercrítico.
Maiores detalhes sobre este sistema coaxial podem ser encontrados no Capítulo 7.
Figura 2.4.2.1.1 - Equipamento comercial para formação de partículas via SAS
(THAR TECHNOLOGIES, 2010)
28
2.4.2.2 Para Formação de Partículas Via RESS
Processos de formação de partículas via RESS são desenvolvidos
principalmente em equipamentos “Home-made”, apesar de existirem comercialmente
equipamentos para o desenvolvimento de tais processos. Isto deve-se principalmente a
simplicidade em instrumentação para o desenvolvimento destes processos. A figura
2.4.2.2.1 apresenta o fluxograma da unidade experimental comercial da Thar Technologies
RESS-100.
Figura 2.4.2.2.1 - Fluxograma da unidade experimental comercial da Thar
Technologies RESS-100. 1 - Reator; 2 - Agitador; 3 – Termostato; 4, 7 e 8 - válvulas; 5 Medidor de fluxo; 6 - Bomba de alta pressão; 9 - Dispositivo de expansão; 10 - Trocador de
calor para aquecimento; 11 - Câmara de expansão; 12 - Trocador de calor para
resfriamento; 13 - Computador (GIL’MUTDINOV et al., 2009).
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35
36
CAPÍTULO 3 - DETALHAMENTO DA CONSTRUÇÃO E FUNCIONAMENTO DA
UNIDADE MULTIPROPÓSITO
A unidade multipropósito para desenvolvimento de processos com fluidos
pressurizados construída pode ser utilizada em laboratórios públicos e privados de centros
de pesquisa, universidades e indústrias para fins de ensino, pesquisa e desenvolvimento de
processos e produtos. A unidade multipropósito possibilita o desenvolvimento e realização
de diferentes processos com fluidos pressurizados, tais como extração com fluidos
pressurizados: extração sub ou supercrítica sem ou com co-solvente, sem ou com
separadores, assistida ou não por ultrassom, extração com líquidos pressurizados; extração
utilizando explosão com CO2 a alta pressão; pasteurização com CO2 a alta pressão; reação
em fluidos pressurizados; formação de micro e nano partículas via RESS (Rapid Expansion
of Supercritical Solutions) com precipitação em condições ambientes ou condições de
pressão e temperatura escolhidas ou via SAS (Supercritical fluid Anti-Solvent), dentre
outros processos.
Nos processos de extração, formação de partículas, dentre outros, geralmente
solventes ambientalmente corretos são empregados, tais como dióxido de carbono, água ou
etanol, porém a referida unidade possibilita a utilização de outros solventes, tais como
isopropanol, entre outros.
A parte estrutural da unidade foi montada utilizando perfilados quadrados de
alumínio de 45 mm (Figura 3.1). Tubulações de aço inox 316 sem costura nos diâmetros de
1/8”e 1/16” foram utilizados para interligar os componentes dos sistemas que suportam
pressões de trabalho de até 800 bar.
37
A unidade contempla componentes fixos e móveis, sendo que os móveis foram
construídos para que qualquer pessoa possa fazer as pequenas modificações com o mínimo
esforço.
Figura 3.1 - Foto da parte estrutural da unidade.
Os componentes fixos são: uma bomba pneumática, uma “válvula Back
Pressure regulator”, uma bomba de HPLC, 2 banhos sendo, um para aquecimento e outro
para resfriamento de alguns componentes da unidade, 4 manômetros e 1 termopar para
medição da pressão e temperatura, respectivamente, em diferentes pontos do sistema, duas
plataformas de tamanhos diferentes construídas para os componentes móveis serem
incorporados, 7 válvulas de bloqueio e uma válvula micrométrica com sistema elétrico de
aquecimento para evitar congelamento e entupimento da linha devido ao efeito JouleThomsom, 1 sistema de aquecimento elétrico tipo jaqueta (para o vaso de pressão menor), 2
38
controladores de temperatura (um para o sistema de aquecimento da válvula micrométrica e
outro para a jaqueta), um rotâmetro e um totalizador de fluxo.
E os componentes móveis são: 1 vaso de pressão de 6,57 mL
(aproximadamente 2 cm de diâmetro e 4,5 cm de altura) com entrada e saída em suas
extremidades, e um banho ultrassônico; 1 vaso de pressão de 500 mL (aproximadamente
6,5 cm de diâmetro e 17 cm de altura) com fundo chato encamisado (possibilita
temperaturas de operação de -10 a 90 ºC por possuir duas serpentinas - uma para fluido
refrigerante do banho de resfriamento e outra para água do banho de aquecimento) com 3
orifícios em sua extremidade superior que, por exemplo, ora funciona como um separador
no processo de extração, possibilitando o fracionamento do extrato obtido, ora funciona
como câmara de precipitação nos processo de formação de partículas via RESS e SAS, e
diferentes acessórios contendo pequenos fragmentos de tubulações previamente construídas
para possibilitar um fácil e rápido rearranjo das linhas da unidade para realizar os diferentes
processos. Maiores informações sobre os procedimentos a serem adotados para a realização
dos processos que a unidade possibilita desenvolver, bem como fluxogramas/esquemas de
cada processo são encontradas no Apêndice IV desta tese.
Em uma planta química tradicional constituída de tubulações, bombas, válvulas,
filtros, vasos de pressão, etc. estes elementos estão dispostos em um arranjo pré-definido e
quaisquer modificações que se deseje realizar na planta necessita da inserção ou retirada de
tubulações, bombas, válvulas, etc. Já na unidade multipropósito, nenhuma bomba, válvula,
vaso de pressão é retirado da unidade. O que ocorre é somente a inserção (“encaixe”) ou
remoção (“desencaixe”) de fragmentos de tubulações seguida de uma posterior limpeza da
tubulação para a adequação da unidade para operar um novo processo.
39
Para a realização de outros processos que também podem ser feitos na referida
unidade multipropósito (Figura 3.2), tais como reação, impregnação, adsorção,
recobrimento de partículas utilizando fluidos pressurizados, dentre outros, é necessário a
escolha correta de qual dos acessórios, bem como de qual dos dois vasos de pressão, ou se
os dois, serão utilizados. Por exemplo, para o desenvolvimento de extrações, somente o
vaso de pressão menor, juntamente com o sistema de aquecimento elétrico e os acessórios
que possibilitem a união das partes móveis com as fixas, devem ser utilizados.
Figura 3.2 - Fotos da unidade multipropósito. A - Vista de frente da unidade; B - Sistema
ultrassônico quando acoplado à unidade; C - Vista do sistema de aquecimento do vasos de
pressão menor: D - Vista lateral da unidade.
40
Do modo como a unidade multipropósito foi projetada ela permite facilmente,
ainda, a sua integração/acoplamento com sistemas de análise (por exemplo, sistemas de
cromatografia), a sua automação por controle via CLP (Controle Lógico Programável) e
IHM (Interface Homem Máquina), assim como a substituição dos seus atuais componentes
móveis (vasos de pressão, por exemplo) por componentes com capacidade diferente. Para
uma ampliação de escala em graus maiores, a unidade possui uma limitação relacionada à
capacidade dos componentes fixos (bombas, banhos, etc.). Porém, com a substituição
destes é possível fazer o escalonamento da unidade para quaisquer dimensões.
Basicamente, a unidade multipropósito pode ser dividida em 4 partes:
Parte 1 - Alimentação do CO2. Esta parte da unidade é composta por bomba
pneumática, uma “válvula Back Pressure regulator”, 1 banho de resfriamento, tubulações e
válvulas de bloqueio. CO2 é pressurizado até altas pressões. Para isto, o solvente é,
primeiramente, refrigerado através da passagem por uma serpentina imersa (extensão de 7
metros) no banho termostatizado de resfriamento (Figura 3.3) para se liquefazer a fim de
ser bombeado/pressurizado pela bomba pneumática.
Parte 2 - Alimentação do solvente líquido. Esta parte da unidade é composta
por bomba de HPLC, tubulações e válvulas de bloqueio. O solvente líquido (H2O, solventes
orgânicos) é pressurizado até altas pressões utilizando uma bomba de HPLC.
Parte 3 - Desenvolvimento dos processos com fluido pressurizado. Esta
parte da unidade é composta por 2 vasos de pressão, banho de aquecimento, banho de
resfriamento, sistema de aquecimento elétrico tipo jaqueta (para o vaso de pressão de 6,57
mL), tubulações, válvulas de bloqueio e válvula micrométrica com sistema de aquecimento.
O fluido pressurizado [puro ou uma mistura (solvente+co-solvente)] executam sua função
41
específica (solvente de extração, solvente ou anti-solvente para formação de partícula,
carbonatação de água, manter uma atmosfera asséptica, proporcionar rompimento celular,
facilitar a extração de metabólitos secundários de fontes vegetais, inchamento do agente
encapsulante, etc.) durante um determinado tempo.
Figura 3.3 - Foto da serpentina imersa no banho termostatizado de resfriamento.
Parte 4 - Medição dos parâmetros de processo. Esta parte da unidade é
composta por medidor de temperatura (1 termopar) e pressão (4 manômetros),
controladores de temperatura (um para o sistema de aquecimento da válvula micrométrica e
outro para a jaqueta), um rotâmetro e um totalizador de fluxo. A medição dos parâmetros:
temperatura e pressão do processo, vazão de CO2 e vazão do solvente líquido são
continuamente medidas durante o desenvolvimento do processo.
42
CAPÍTULO 4 - ANTIOXIDANT PIGMENT EXTRACTION USING A HOMEMADE PRESSURIZED SOLVENT EXTRACTION SYSTEM
Diego T. Santos and M. Angela A. Meireles
Trabalho submetido ao periódico International Journal of Food Engineering.
43
Key words
Antioxidant pigments, Pressurized Solvent Extraction, Anthocyanins, Bixin, Home-made
equipment
Abstract
Increasing reports on health hazards and toxicity of synthetic pigments are driving the food
industry towards application of natural colorants in an increasing number of processed food
products. The attention that natural dyes is getting is also due to the functional properties
attributed to some of these pigments. Commonly, conventional extraction methods are used
to extract these compounds from natural sources, nevertheless, these methods are, in
general, time and solvent consuming and may promote the degradation of these
compounds. In order to overcome these drawbacks short time extraction conditions using
environmentally friendly pressurized solvents, such as supercritical carbon dioxide and
pressurized ethanol have been used successfully to obtain antioxidant pigments-rich
extracts. The objective of this work was to validate a home-made pressurized solvent
extraction system that can be used for Supercritical Fluid Extraction (SFE) and Pressurized
Liquid Extraction (PLE) processes, independently. For this, functional pigment sources
such as Annatto seeds and Jabuticaba skins were used as model plant materials. The
feasibility of the integration of an ultrasonic system to our home-made unit was also
evaluated. It was demonstrated that our home-made pressurized solvent extraction
consistently leaded to the expected results with good reproducibility. Furthermore, it was
shown that one step (SFE or PLE) or two steps extraction can be effectively carried out
using this apparatus.
4.1 Introduction
Increasing reports of health hazards and toxicity of synthetic pigments are
driving the food industry towards application of natural colorants in an increasing number
44
of processed food products. Commonly, conventional extraction methods are used to
extract these compounds from natural sources, nevertheless, these methods are, in general,
time and solvent consuming and may promote the degradation of these compounds.
In order to overcome these drawbacks short time extraction conditions using
pressurized solvent methods, such as Supercritical Fluid Extraction (SFE) and Pressurized
Liquid Extraction (PLE) methods have been used successfully to obtain antioxidant
pigments-rich extracts [1, 2].
SFE has important advantages over traditional extraction techniques, mainly
considering that low volumes of solvents, if any, are employed and that a solvent free
extract can be obtained [3]. On the other hand, the use of PLE employing GRAS (Generally
Recognized as Safe) such as ethanol and water as solvent allows the attainment of generally
higher extraction yields also limiting the use of toxic organic solvents [4]. Moreover, high
pressure extraction has other advantages that should be considered, such as the fact that
native enzymes, which degrade some compounds, are inhibited by extraction pressure
increasing and/or CO2 addition [5].
Carotenoids and anthocyanins are two of the most widely used dyes in food,
pharmaceutical and cosmetic industries. Carotenoids are a diverse group of lipophilic
compounds that contribute to the yellow to red colors of many foods. They are polyenes
consisting of 3 to 13 conjugated double bonds and in some cases 6 carbon ring structures at
one or both ends of the molecule [6]. Anthocyanins belong to the phenolic compound class
forming an important class of natural pigments found in flowers, fruits, berries, among
others. Anthocyanins can be useful as colorants (red and blue colors), and/or for human
health, as they are antioxidants and free radical scavengers [7].
45
Annatto seed extracts are orange/red natural carotenoid colouring agents
obtained from the outer coats of the seeds of Bixa orellana L. The Annatto (Bixa orellana
L.) is the fruit from “urucuzeiro”, native plant from the tropical America, also cultivated in
Asia and Africa. Although it is produced in many tropical countries, Peru and Brazil are the
largest producers of this seed. The major pigment bixin, present only in this plant, is a red
carotenoid with high socio-economic potential and is extensively used in textile,
pharmaceutical, cosmetic and food industries. Carotenoids of Annatto seeds have proven
antioxidant activity against free radicals and also, are capable of blocking the sunlight [8].
Jabuticaba skins extracts are purple natural anthocyanin colouring agents
obtained from skins of Myrciaria cauliflora. The Jabuticaba (Myrciaria cauliflora) is the
fruit from Jabuticabeira, a tree that grows mainly in Brazil, most frequently in the states of
São Paulo, Minas Gerais, Rio de Janeiro, and Espírito Santo. Jabuticaba is grape-like in
appearance and texture, although its skin is thicker and tougher. This fruit has a dark purple
to almost black skin color due to a high content of anthocyanins that covers a white
gelatinous flesh inside. It is 3 to 4 cm in diameter and carries from one to four large seeds.
The fruits are born directly on the main trunks and branches of the plant, lending to a
distinctive appearance to the fruiting tree [7]. Anthocyanins of Jabuticaba skins obtained by
different extraction methods have proven antioxidant activity [9].
In this work a home-made pressurized solvent extraction system was designed
and built, in which pure supercritical CO2 (SFE) and GRAS solvent (PLE) can be used
independently. SFE was used for obtaining Annatto seed extract and PLE for Jabuticaba
skin extract. Home-made SFE results were compared to those obtained using a commercial
SFE unit using the same processing conditions. The feasibility of the use of an ultrasonic
46
system to assist supercritical CO2 extraction of carotenoid pigments from Annatto seeds
using our home-made system was also studied. Moreover, fractionated extractions of
Jabuticaba skins were performed in two steps: i) a first step (PLE), wherein pressurized
ethanol was used in order to extract polar compounds like anthocyanin pigments; ii) a
second step, wherein supercritical CO2 was used in order to recovery low polarity CO2soluble compounds. The chemical compositions of both extracts were determined.
4.2 Materials and methods
4.2.1 Plant Material
4.2.1 Annatto seeds
Annatto seeds (Bixa orellana L.) cultivated in the state of São Paulo (variety
Piave) were supplied by Agronomical Institute of Campinas (IAC). The humidity of the
dried skins was determined by the AOAC method [10]. The seeds at 12.27 % moisture
were stored in the dark in a domestic freezer (-10 ºC) (Double Action, Metalfrio, São Paulo,
Brazil) until extraction.
4.2.2 Jabuticaba skins
Jabuticaba fruits (Myrciaria cauliflora) harvested from a plantation in the State
of São Paulo, Brazil, were acquired from a fruit and vegetable market centre (CEASACampinas, Brazil). Immediately after acquired, the fruits were manually peeled and the
Jabuticaba skins were dried for a few hours at 45 ºC in an oven with forced air circulation
(Marconi, MA 035/1, Piracicaba, São Paulo, Brazil). The humidity of the dried skins was
determined by the AOAC method [10]. The dried skins at 66.52 % moisture were cut into
47
approximately 5 mm cubes and stored in the dark in a domestic freezer (-10 ºC) (Double
Action, Metalfrio, São Paulo, Brazil) until extraction.
4.2.2 Extraction Procedures
A schematic diagram of the home-made pressurized solvent extraction system
is given in Figure 4.2.2.1. The equipment consists of a thermostatic bath (Marconi, MA184, Piracicaba, Brazil), an air driven liquid (CO2 pump) (Maximator Gmbh, PP 111,
Zorge, Germany), a HPLC pump (Thermoseparation Products, Model ConstaMetric 3200
P/F, Fremoni, USA) and a stainless steel extraction cell (6.57 mL, Thar Designs, Pittsburg,
USA) containing syntherized metal filters at the inlet and outlet.
3
4
2
7
3
3
4
5
4
4
6
12
10
1
4
4
11
8
13
9
Figure 4.2.2.1 - Schematic diagram of the home-made pressurized solvent extraction
system. 1 CO2 cylinder; 2 CO2 filter; 3 Manometers; 4 Valves; 5 Thermostatic bath; 6 CO2
Pump; 7 Back pressure regulator; 8 HPLC pump; 9 Solvent resevoir; 10 Extraction cell; 11
Micrometric valve with a heating system; 12 Temperature controller; 13 Sampling bottle.
48
4.51 g of plant material (whole Annatto seeds or dried/cut Jabuticaba skins) was
first loaded into the extraction cell, filled with extraction solvent (CO2 or ethanol) and then
pressurized. During 5 min the sample was heated to ensure that the extraction cell would be
at the desired temperature during filling and pressurization procedures. After pressurization,
the sample with pressurized solvent was kept statically at the desired pressure for a desired
time (static extraction time). Thereafter, the blocking and micrometric valves were
carefully opened keeping the pressure constant to the desired flow rate being the extraction
cell rinsed with fresh extraction solvent during a certain time (dynamic extraction time).
For SFE process, liquid CO2 99.9 % (Gama Gases Especiais Ltda., Campinas,
Brazil) was fed from the cylinder through a thermostatic bath at -10 °C to ensure the
liquefaction of the fluid and to prevent cavitation, and then it was pumped by the CO2
pump to the extraction cell containing Annatto seeds or Jabuticaba skins previously
extracted with pressurized ethanol. The SFE yields (the total amount of extractable solute)
were calculated as the ratio between the total extract mass and the feed mass (dry basis).
Using Annatto seeds the processing conditions were: Pressure 31 MPa,
Temperature 333 K, Static extraction time 10 min and CO2 flow rate 1.74.10-4 kg/s during
240 min. Experiments also using Annatto seeds at the same processing conditions were
performed in a commercial SFE system (Spe-ed SFE unit, Applied Separations, model
7071, Allentown, USA) and in the home-made system (Figure 4.2.2.2) working with the
extraction cell immerserd into a Unique Maxi Clean 1400 40 kHz (135 W) ultrasonic water
bath (Indaiatuba, Brazil) with or without ultrasound irradiation during the dynamic
extraction time. To keep the ultrasonic water bath at a fixed temperature it was used a
heating water bath with external recycling (Marconi, MA 127BO, Piracicaba, Brazil). The
49
tubing line was washed with ethanol after extraction to recover the extract deposited on it in
order to calculate the percentage of residual extract (%). The total extract mass was
determined by the sum of the extract obtained during the extraction and the extract
recovered in the cleaning process. The extraction pressure and temperature used were
chosen to compare our results with those of Chao et al. [17] study.
3
4
2
7
3
3
4
14
11
5
4
4
10
6
4
13
1
4
12
15
8
9
Figure 4.2.2.2 - Schematic diagram of the home-made pressurized solvent extraction
system. 1 CO2 cylinder; 2 CO2 filter; 3 Manometers; 4 Valves; 5 Thermostatic bath; 6 CO2
Pump; 7 Back pressure regulator; 8 HPLC pump; 9 Solvent reservoir; 10 Extraction cell; 11
Ultrasound bath; 12 Heating bath; 13 Micrometric valve with a heating system; 14
Temperature controller; 15 Sampling bottle.
For Jabuticaba skins previously extracted with pressurized ethanol the
processing conditions were: Pressure 300 bar, Temperature 60 ºC, Static extraction time 5
50
min and CO2 flow rate 1.67.10-4 kg/s (during 20 min). The extraction pressure and
temperature employed in this work were selected based on our previous experiments.
For PLE process, ethanol 99.5 % (Ecibra, Santo Amaro, Brazil) was pumped
directly to the extraction cell containing Jabuticaba skins by the HPLC pump. Based on our
previous experiments, extraction pressure, temperature, static extraction time and solvent
flow rate were set at 50 bar, 80 ºC, 9 min and 1.67 mL/min (during 12 min), respectively.
After PLE, anthocyanin extracts were rapidly cooled to 5 °C in ice water to prevent
anthocyanin degradation. Subsequently, the extraction cell was exaustively purged with a
flow rate of 0.71 kg/h of carbon dioxide 99.9% (Gama Gases Especiais Ltda., Campinas,
Brazil) during 8-9 min to ensure that no residual anthocyanin extract solution would be into
the extraction cell. At the end, ethanol from the extract solution was evaporated using a
rotary evaporator (Laborota, model 4001, Vertrieb, Germany), with vacuum control
(Heidolph Instruments Gmbh, Vertrieb, Germany) and thermostatic bath at 40 °C. All the
extracts were stored (- 10 ºC) in the dark until analysis.
All extractions were done in duplicate.
4.2.3 Extract Characterization
4.2.3.1 From Jabuticaba skins
4.2.3.1.1 Anthocyanin content
The anthocyanin content of the Jabuticaba skin extract obtained by PLE and
SFE method was determined using the pH differential method exactly as described by
Santos et al. [9], which relies on the structural transformation of the anthocyanin
chromophore as a function of pH.
51
4.2.3.1.2 Thin-Layer Chromatography (TLC)
The supercritical extracts from Jabuticaba skins previously extracted with
pressurized ethanol were fractionated by thin-layer chromatography (TLC). The TLC was
performed using silica plates (20 x 20 cm, 1-mm height, Merck, Darmstadt, Germany) and
four different sprays to reveal the compounds present in the extracts. The mobile phase
used was composed by hexane 70 % v/v (96 %, P.A., Merck, Darmstadt, Germany) and
ethyl acetate 30 % v/v (99.5 %, P.A., Merck, Darmstadt, Germany). 2-Diphenylpicrylhydrazyl radical (DPPH) a purple-colored stable free radical is reduced into the
yellow colored diphenylpicryl hydrazine. The test was performed with a rapid TLC
screening method using a 0.2 % (v/v) DPPH in Methanol. Thirty minutes after spraying
active compounds, if the extract has antioxidant activity, yellow spots appear against purple
background. To observe compounds as flavonoids on the ultraviolet light (365 nm) the
spray solution of 2-aminoethyl diphenylborinate (Sigma, lot 123k2512, U.S.A.) was 1 % in
methanol. To observe the compounds of the volatile oil on the visible and ultraviolet (365
nm) light spray of anisaldehyde solution was used. Finally, to observe alkaloids on
ultraviolet (254 nm) light no treatment was done and to observe alkaloids on visible light
the spray solution of Dragendorff was used [11].
4.2.4 Statistical Analysis
For establishing the statistical significant differences or similarities between the
values of extraction yields obtained using different SFE systems/configurations, the
Tukey’s test was used. A confidence coefficient of 95 % was used for the comparison of all
the mean’s pairs.
52
4.3 Results and discussion
4.3.1 Obtaining Annatto seed extracts
SFE was used for obtaining Annatto seed extracts. Home-made SFE results
were compared to those obtained using a commercial SFE using the same processing
conditions. The feasibility of the use of an ultrasonic system to assist supercritical CO2
extraction of carotenoid pigments from Annatto seeds using our home-made apparatus was
also studied.
Figure 4.3.1.1 shows that the behavior of the extraction curves was very similar,
independently of the system/configuration used. Nobre et al. [12] using a similar SFE
apparatus observed analogous extraction curves for pigment extraction using also pure
supercritical CO2 and whole Annatto seeds.
On the other hand, it seems that the recoveries of pigments obtained were low
and decreased with the use of our home-made system (Figure 4.3.1.1). Moreover, the
recoveries were even lower with the use of ultrasonic system to assist supercritical CO2
extraction. In order to verify if the ultrasonic irradiation has affected negatively the
extraction process a control experiment was done, using the ultrasonic system as only a
water bath heating for the extraction cell. It was demonstrated that the recovery of the
pigments was not affected by the ultrasonic irradiation. However, apparently the use of a
different system to heat the extraction cell leaded to recovery decrease.
53
4
3.5
Yield (%) d.b.
3
2.5
2
1.5
1
0.5
0
0
40
80
120
Time (min)
160
200
240
Figure 4.3.1.1 - Recovery of pigments from Annatto seeds as a function of time using
different SFE systems/configurations: ■ Commercial SFE; □ Home-made SFE using
electric heating; ● Home-made Ultrasound Assisted SFE; ○ Home-made SFE using water
bath heating.
When the extraction yields were calculated after 240 min summing the extract
obtained during the extractions with the extract recovered in the cleaning of the tubing line
after the extraction cell, it was confirmed that the recovery decrease using the home-made
system, independently of the configuration, was due to a large amount of residual extract
depositated in the exit tubing line and not to the SFE system/configuration employed
(Figure 4.3.1.2). Table 4.3.1.1 shows that a significant higher amount of residual extract
deposited in the tubing line when using our home-made system, independently of the
configuration, compared to those obtained using the commercial SFE apparatus.
54
Home-made SFE
using water bath
heating
Home-made
Ultrasound Assisted
SFE
Home-made SFE
using electric heating
Comercial SFE
0
0,5
1
1,5
2
2,5
3
3,5
4
Yield (%)
Figure 4.3.1.2 - Extraction yields (%) (after 240 min) using different SFE
systems/configurations.
Comparing the extraction yield values (after 240 min) obtained using our homemade system, independently of the configuration, to those obtained using the commercial
SFE apparatus at the same processing conditions, it was verified by Tukey’s test that their
differences were not significant, indicating that statistically our home-made system may
produce equal extraction yields with similar percentage standard deviation (Table 4.3.1.1),
demonstrating that this system also produces experimental data with good reproducibility.
Additionally, it was concluded that the use of our ultrasonic system to enhance supercritical
CO2 extraction of antioxidant pigments from Annatto seeds have not influenced the
extraction process.
Table 4.3.1.1 - Percentage of residual extract of total extract (%) deposited in the exit
tubing line when using different SFE systems/configurations.
Residual Extract (%)
Commercial SFE
7.17 ± 1.11
Home-made SFE using electric heating
32.95 ± 4.80
Home-made Ultrasound Assisted SFE
35.24 ± 2.59
Home-made SFE using water bath heating
37.74 ± 5.49
55
In contrast to our findings, some authors have demonstrated that SFE assisted
by ultrasound give higher extraction yields than SFE without ultrasound irradiation for
extracting diverse bioactive compounds [13-15]. A possible explanation to our results may
be related to the low ultrasonic power of our system. Balachandran et al. [13] using similar
ultrasonic system (ultrasonic transducer placed outside the extraction cell and inside a water
bath) with a 3.7-fold higher ultrasonic power than that used in the present work obtained a
significant increase in the extraction process of pungent compounds from ginger sample.
Other studies using similar ultrasonic power or even lower than that used in our study
succesfully resulted in better extraction yields through integrating the ultrasonic transducer
inside the supercritical extractor or in the upper part of the extraction cell [14]. Thus, to
have an effective positive influence of the ultrasound irradiation during our SFE processes
or the transducer power should be increased or the transducer should be placed inside the
extractor.
Comparing the results of this study to literature data we observed that the yield
differences might be mainly associated to the effects of the origin, year of production, preprocessing storage conditions, treatments of the samples, among others [16]. Chao et al.
[17] obtained at the same extraction pressure and temperature and similar CO2 flow rate
and extraction dynamic time approximately 1.47-fold lower extraction yields than those
obtained in this study.
4.3.2 Obtaining Jabuticaba skin extracts
Fractionated extractions of Jabuticaba skins were performed in two steps: i) a
first step (PLE), wherein pressurized ethanol was used in order to extract polar compounds
like anthocyanin pigments; ii) a second step, wherein supercritical CO2 was used in order to
recover low polarity CO2-soluble compounds.
The chemical composition the PLE extract was characterized to the presence of
anthocyanins. Visually the extract solution presented a purple color. The anthocyanin
content of the PLE extract was 2.49 ± 0.54 mg cyanidin-3-glucoside/g dry material.
Otherwise, during the re-extraction of the Jabuticaba skins previously extracted by PLE
56
using supercritical CO2 an extract presenting a yellow-green color, with insignificant
anthocyanin content, was obtained at the processing conditions employed.
Previously we have verified that these yellow-green extracts present antioxidant
activity and are only extracted from Jabuticaba skins using pure CO2. Given that SFE
experiments using ethanol as co-solvent resulted in a purple extract (rich in anthocyanins)
(results not shown), in order to obtain only the yellow-green extract after the first step, CO2
was purged into the extraction cell to eliminate almost all residual ethanol from the
vegetable matrix that could act as co-solvent.
The SFE extraction yield employing pressure and temperature of 30 MPa and
333 K, respectively, was 1.02 ± 0.15 % and the SFE extracts (second step) were
characterized by thin-layer chromatography (TLC).
Figure 4.3.2.1 shows that both extracts present similar phytochemical profile in
the TLC plates, indicating that this fractionated extraction process is reproducible. Indeed,
it might be suggested that the designed home-made system can produce reliable results also
to this combined extraction process.
In Figure 4.3.2.1-a is shown that the SFE extract presents antioxidant activity.
Flavonoid compounds have characteristics to form blue fluorescent zones in
UV-365 nm. Natural products (NP) reagent revealed blue fluorescence zones in SFE
Jabuticaba skin extracts near the solvent front which can be seen in figure 4.3.2.1-b.
According to Wagner and Bladt [11], depending on the structural type, flavonoids show
dark yellow, green or blue fluorescence. Since a blue fluorescence has similar Rf values to
the yellow spots of the DPPH plate, the antioxidant activity of this extract could be related
to the flavonoid content.
Essential oils are volatile, odorous principles consisting of terpenes alcohols,
aldehydes, ketones and esters (> 90 %) and/or phenylpropane derivatives. The
anisaldehide-sulphuric acid reagent revealed (Figure 4.3.2.1-c) two green-violet zones that
can be terpenoids, propylpropanoids, pungent and bitter principles and saponins due their
coloration characteristics on visible light. According to Wagner and Bladt [11], most of
these compounds develop fluorescence under UV-365, which was observed for these
57
extracts (Figure 4.3.2.1-d). Since one of the green-violet zones is placed at a similar length
in the TLC place of the yellow spots of the DPPH plate and supercritical CO2 is a good
solvent for essential oils extraction [3], probably the antioxidant activity of this extract is
associated to the essential oil content.The presence of alkaloids was not detected after
treatment with Dragendorff reagent for the Jabuticaba skin extract obtained by SFE.
Seabra et al. [5] also observed these yellow-green extracts using elderberry
pomace as anthocyanin source using supercritical CO2. It was suggested in their
experiments, that these yellow-green extract are rich in phenolic compounds although no
antioxidant activity evaluation and chemical characterization were done.
Figure 4.3.2.1 - Thin-layer chromatography (TLC) plates of jabuticaba extracts obtained
using supercritical CO2 (1 - Extract obtained at 30 MPa/333 K run 1; 2 – Extract obtained
at 30 MPa/333 K run 2; a) Revealed using DPPH reagent on visible light; b) Revealed
using Natural products (NP) reagent on light (365 nm); c) Revealed using anisaldehidesulphuric acid reagent on visible light; d) Revealed using anisaldehide-sulphuric acid
reagent on ultraviolet light (365 nm).
58
4.4 Conclusions
In this work we validated a home-made pressurized solvent extraction system
that can be used for Supercritical Fluid Extraction (SFE) and Pressurized Liquid Extraction
(PLE) processes, independently, using Annatto seed and Jabuticaba skin as model plant
materials.
The feasibility of the integration of a ultrasonic system to our home-made unit
was not succesfully achieved, though with some small modifications Ultrasound-Assisted
supercritical Fluid Extraction (USFE) could be carried out using this home-made apparatus.
Fractionated extractions of Jabuticaba skins were successfully performed using
our apparatus, producing two valuable extracts with antioxidant activities: one rich in
anthocyanin pigments; another rich in lipophilic compounds that can be essential oil and
less polar flavonoid compounds.
Acknowledgements
The authors are grateful to CNPq for financial support. Diego T. Santos thanks
CNPq (141894/2009-1) for the doctorate fellowship.
References
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25 (2008), 419-426.
[2] P. Arapitsas, C. Turner. Talanta, 74 (2008) 1218-1223.
[3] D. T. Santos, M. A. A. Meireles. Recent Patents on Eng., In press (2010).
[4] Z. Y.Ju, L. R. Howard. J. Agric. Food Chem., 51 (2003) 5207-5213.
[5] I. J. Seabra, M. E. M. Braga, M. T. Batista, H. C. Sousa. J. Supercrit. Fluids, 54 (2010)
145-152.
[6] D. J. McClements, E. A. Decker, Y. Park, J. Weiss. Crit. Rev. Food Sci. Nutrition, 49
(2009) 577-606.
[7] D. T. Santos, M. A. A. Meireles. Pharmacog. Rev., 3 (2009) 127-132.
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[8] C. L. C. Albuquerque, M. A. A. Meireles. Estimate of the Cost of Manufacturing
(COM) of Natural Colorants Obtained by Supercritical Fluid Extraction. II Iberoamerican
Conference on Supercritical Fluids, Natal, Brazil (2010).
[9] D. T. Santos, P. C Veggi, M. A. A. Meireles. J. of Food Eng., 101 (2010) 23-31.
[10] AOAC (Association of Official Analytical Chemists) Official analysis. 16ª ed. 3ª rev.
Washington, USA (1997).
[11] H. Wagner, S. Bladt, S., Plant Drug Analysis: A Thin Layer Chromatography Atlas, 2ª
ed., Springer-Verlag Berlin Heidelbeg, New York, USA (2001).
[12] B. P. Nobre, R. L. Mendes, E. M. Queiroz, F. L. P. Pessoa, J. P. Coelho, A. F. Palavra.
Brazilian J. Chem. Eng., 23 (2006) 251-258.
[13] S. Balachandran, S. E. Kentish, R. Mawson, M. Ashokkumar. Ultras. Sonochem., 13
(2006) 471-479.
[14] E. Riera, A. Blanco, J. García, J. Benedito, A. Mulet, J. A. Gallego-Juárez, M. Blasco.
Ultrasonics, 50 (2010) 306-309.
[15] A. Hu, S. Zhao, H. Liang, T. Qiu, G. Chen. Ultras. Sonochem., 14 (2007), 219-224.
[16] M. Al-Farsi, C. Alasalvar, A. Morris, M. Baron, F. Shahidi. J. Agric. Food Chem., 53
(2005) 7592-7599.
[17] R. R. Chao, S. J. Mulvaney, D. R. Sanson, F. Hsieh, M. S. Tempesta. J. Food Sci., 56
(1991) 80-83.
60
CAPÍTULO 5 - PRESSURIZED LIQUID EXTRACTION OF PHENOLIC
COMPOUNDS FROM JABUTICABA SKINS: OPTIMIZATION STUDY
Diego T. Santos and M. Angela A. Meireles
Trabalho submetido ao periódico Journal of Food Engineering.
61
Key words
Anthocyanins, Phenolic compounds, Pressurized Liquid Extraction, Jabuticaba skins,
Myrciaria cauliflora
Abstract
Conventional extraction methods are normally used for anthocyanin extraction,
nevertheless, these methods are time and solvent consuming and may promote the
degradation of these compounds. In order to overcome these drawbacks high-temperatureshort time extraction conditions have been used successfully to obtain an anthocyanin-rich
extract. Optimization of the extraction of anthocyanins and other phenolic compounds from
jabuticaba skins, a promising Brazilian source of these compounds, was studied using
Pressurized Liquid Extraction. Optimization study was carried out using ethanol as solvent,
extraction pressure (50-100 bar), temperature (40-120 ºC) and static extraction time (3-15
min) as independent variables. The optimum PLE conditions for all response variables were
estimated, however specific PLE conditions (highest recovery of anthocyanins) were
chosen for comparison with a conventional Low Pressure Liquid Extraction (LPLE)
method in terms of yield, composition (anthocyanins and phenolic compounds), and
economical feasibility. Response surface methodology successfully optimized and modeled
our extraction process.
5.1 Introduction
Anthocyanins belong to the phenolic compound class forming an important
class of natural pigments found in flowers, fruits, berries, among others. Anthocyanins can
62
be useful as colorants (red and blue colors), and/or for human health, as they are
antioxidants and free radical scavengers (Santos and Meireles, 2009).
Conventional analytical anthocyanin extraction methods depend on solid-liquid
extraction, where organic solvents such as methanol, ethanol or acetone are normally used.
However, classical extraction methods are time and solvent consuming and may promote
the anthocyanin degradation during the extraction process (Santos et al., 2010).
Elevated temperatures are reported to improve the efficiency of extraction due
to enhanced diffusion rate and solubility of analytes in solvents. Nevertheless, elevated
extraction temperatures may simultaneously increase the rate of anthocyanin degradation.
Conventional extraction and purification/concentration of anthocyanins is typically
conducted at temperatures ranging from 20 to 50 °C, because temperatures > 70 °C have
been shown to cause rapid anthocyanin degradation (Ju and Howard, 2003).
As the degradation rate of anthocyanins is time and temperature dependent,
high-temperature-short time extraction conditions have been used successfully to obtain an
anthocyanin-rich extract (Gizir et al., 2008).
The use of Pressurized Liquid Extraction (PLE) technique is an attractive
alternative, since it allows fast extraction and small solvent consumption. Sometimes
referred to as Pressurized Solvent Extraction (PSE®) and Accelerated Solvent Extraction
(ASE®), PLE has been successfully used for extraction of thermolabile anthocyanins from
various plants (Petersson et al., 2010).
PLE enables rapid extraction (less than 30 min) of analytes in a closed and inert
environment, under high pressures (no higher than 200 bar) and temperatures (room
temperature-200 °C). A major advantage of PLE over conventional solvent extraction
63
methods conducted at atmospheric pressure is that pressurized solvents remain in a liquid
state well above their boiling points, allowing for high-temperature extraction. These
conditions improve analyte solubilities and the desorption kinetics from the matrices
(Richter et al., 1997). Hence, extraction solvents including ethanol and water that are
inefficient in extracting phytochemicals at low temperatures may be much more efficient at
elevated PLE temperatures (Ju and Howard, 2003).
The increasing interest in the health-beneficial properties of phenolic
compounds, such as anthocyanins, has prompted researchers to screen plants for phenolic
content and antioxidant capacity. Recently, it has been reported by our research group that
jabuticaba (Myrciaria cauliflora) skins seem to be a promising source of antioxidant
compounds including anthocyanins. Jabuticaba fruit has a dark purple to almost black skin
that covers a white gelatinous flesh inside (Santos et al., 2010).
This research was undertaken to evaluate PLE of anthocyanins and other
phenolic compounds from the skins of a highly pigmented Brazilian fruit, called jabuticaba,
(Santos and Meireles, 2009) and to investigate how extraction pressure, temperature and
static extraction time affect PLE process using ethanol as solvent. PLE process was
optimized by using Response Surface Methodology (RSM) for the extraction yield,
anthocyanin and total phenolic compounds extraction. Specific PLE conditions (highest
recovery of anthocyanins) were chosen for comparison with a conventional Low Pressure
Liquid Extraction (LPLE) method in terms of extraction yield and extract composition
(anthocyanins and phenolic compounds).
64
5.2 Material and methods
5.2.1 Plant Material
Jabuticaba fruits (Myrciaria cauliflora) harvested from a plantation in the State
of São Paulo, Brazil, were acquired from a fruit and vegetable market centre (CEASACampinas, Brazil). Immediately after acquiring, the fruits were manually peeled and the
jabuticaba skins were dried for a few hours at 45 ºC in an oven with forced air circulation
(Marconi, MA 035/1, Piracicaba, São Paulo, Brazil). The dried skins (66.52 % moisture)
were cut into approximately 5 mm cubes and stored in the dark in a domestic freezer (-10
ºC) (Double Action, Metalfrio, São Paulo, Brazil) until extraction.
5.2.2 Extraction Procedures
5.2.2.1 Pressurized Liquid Extraction (PLE)
The pressurized liquid extraction set up is given in Figure 5.2.2.1. The solvent
was pumped by a HPLC pump (Thermoseparation Products, Model ConstaMetric 3200
P/F, Fremoni, USA) into the extraction cell placed in an electrical heating jacket at the
desired temperature until the required pressure was obtained. All connections within the
system were made of stainless steel tubes (1/16” and 1/8”).
After PLE, anthocyanin extracts were rapidly cooled to 5 °C in ice water to
prevent anthocyanin degradation. Subsequently, the extraction cell was exaustively purged
with a flow rate of 0.71 kg/h of carbon dioxide 99.9% (Gama Gases Especiais Ltda.,
Campinas, Brazil) during 8-9 min to ensure that no residual anthocyanin extract solution
would be into the extraction cell. At the end, ethanol 99.5 % (Ecibra, Santo Amaro, Brazil)
from the extract solution was evaporated using a rotary evaporator (Laborota, model 4001,
Vertrieb, Germany), with vacuum control (Heidolph Instruments Gmbh, Vertrieb,
65
Germany) and thermostatic bath at 40 °C. All the extracts were stored (- 10 ºC) in the dark
until being used in the encapsulation processes.
Solvent reservoir
HPLC Pump
CO2 reservoir
Extractor
Column
Blocking valve
Micrometric valve
Sampling bottle
Figure 5.2.2.1 - Pressurized liquid extraction set-up.
To determine the effects of extraction pressure, temperature and static
extraction time on extraction yield, recovery of anthocyanins and phenolic compounds, the
13 experiments listed in Table 5.2.2.1 were performed as described above. All the
extractions were performed in duplicate. Experiments 9 and 13, central points of the 23 and
complementary 22 full factorial designs, respectively, were done in triplicate.
Table 5.2.2.1 - The experimental design of phenolic compound extraction from jabuticaba
skins
Experiment
Pressure (bar)
Temperature (ºC)
Static extraction time (min)
1
50
40
3
2
100
40
3
3
50
80
3
4
100
80
3
66
5
50
40
9
6
100
40
9
7
50
80
9
8
100
80
9
9
75
60
6
10
50
80
15
11
50
120
9
12
50
120
15
13
50
100
12
5.2.2.2 Conventional Low Pressure Liquid Extraction (LPLE)
Conventional solid-liquid extraction was performed in the percolation regime at
room temperature (22-23 ºC), with the solvent/solution pumped continuously through the
biomass to increase the efficiency of intraparticle mass transfer. 10 g of dried jabuticaba
skin pieces were packed in a bed column and 100 mL of ethanol 99.5 % (Ecibra, Santo
Amaro, Brazil)/solution (solvent plus extract) passed through the packed bed slowly
making sure that the plant material to be extracted was always covered with the remaining
extraction solvent (flow rate of 28.02 cm3/min) towards the bottom of the column under
gravity for 2 hours. After extraction, the solvent was evaporated and the extract was stored,
exactly as described before.
5.2.3 Extract Characterization
5.2.3.1 Anthocyanin content
The Total Monomeric Anthocyanin (TMA) content was determined using the
pH differential method described by Giusti and Wrolstad (2001), which relies on the
67
structural transformation of the anthocyanin chromophore as a function of pH. A UV–Vis
spectrophotometer (Hitachi, model U-3010, Tokyo, Japan) was used for spectral
measurements at maximum absorbance wavelength (approximately 512 nm) and 700 nm,
using distilled water as blank. For this purpose, 20 mg of extract were dissolved in 10 cm3
of distilled water. Two dilutions of the sample were prepared: one with hydrochloric
acid/potassium chloride buffer pH = 1.0 and the other with sodium acetate/acetic acid
buffer pH = 4.5. The pH values of the buffers were measured using a pH-meter (Digimed,
model DM-22, São Paulo, Brazil) calibrated with buffers at pH 4.01 and 6.86 and they were
adjusted with HCl (99.5 % Ecibra, Santo Amaro, Brazil). Aliquots of extract were brought
to pH 1.0 and 4.5; 15 min later, the absorbance of each equilibrated solution was measured
at the maximum absorption wavelength and 700 nm for haze correction using a 1 cm path
length glass cells (l). The dilution factor (DF) was determined (final volume per original
sample volume). The difference in absorbance values at pH 1.0 and 4.5 is directly
proportional to the TMA concentration. The anthocyanin content was calculated as
cyanidin-3-glucoside (MW = 449.2 g/mol and ε = 26.900 L/mol.cm) and the results were
expressed as mg Cy-3-glucoside/g dry material. The absorbance of the diluted sample (A)
and the TMA were calculated with Equations (1) and (2):
A = ( A max − A 700 ) pH 1, 0 − ( A max − A 700 ) pH 4,5
TMA ( mg / L ) = ( A x MW x DF x 1000 ) / (ε x l )
(1)
(2)
5.2.3.2 Total Phenolic Compounds content
Total phenolic content was estimated using the Folin-Ciocalteau method for
total phenolics, based on a colorimetric oxidation/reduction reaction of phenols (Singleton
and Rossi, 1965). Briefly, 1 cm3 of sample (1 mg of extract/1 cm3) was mixed with 1 cm3
68
of Folin and Ciocalteu’s phenol reagent. After 3 min, 1 cm3 of saturated sodium carbonate
solution (50 % w/w) was added to the mixture and the volume adjusted to 10 cm3 with
distilled water. The reaction was kept in the dark for 90 min at room temperature, after
which the absorbance was measured at 725 nm with a UV–Vis Spectrophotometer Hitachi
model U-3010 (Tokyo, Japan). For control sample, 1 cm3 of distilled water was taken. The
results were calculated on the basis of the calibration curve of gallic acid (GA) and
expressed as milligrams of gallic acid equivalents (GAEs)/g dry material.
5.2.4 Statistical Analysis
Statistical analyses were performed using Pareto analysis with consequent
Experimental Design. ‘Statistica’ software (release 7, StatSoft, Tulsa, USA) was first used
to calculate the effects of the extraction conditions (pressure, temperature and static
extraction time) by a 23 full factorial design on extraction yield, on the recovery of
anthocyanins and phenolic compounds employing pressure, temperature and static
extraction time ranges of 50-100 bar, 40-80 ºC and 3-9 min, respectively (experiments 1-9
in Table 5.2.2.1). As the optimum extraction conditions were not achieved a
complementary 22 full factorial design at constant pressure (50 bar) employing larger
temperature (80-120 ºC) and static extraction time (9-15 min) ranges (experiments 10-13 in
Table 5.2.2.1) was carried out. The selected optimum extraction process conditions were
estimated through three dimensional response surface plots of the independent variables
and each dependent variable. Response Surface Methodology (RSM) analysis was also
applied on the data for prediction of optimum conditions of PLE for extraction yield,
anthocyanins and total phenols from jabuticaba skins.
69
5.3 Results and discussion
5.3.1 Effects of process variables on the extraction yield
The effects of extraction pressure, temperature and static extraction time on the
extraction yield were evaluated. At a variable range of 50-100 bar, 40-80 ºC and 3-9 min
this response variable was significantly (95 % confidence level, p<0.05) affected only by
the extraction temperature. On the other hand, at a variable range of 80-120 ºC and 9-15
min at a fixed extraction pressure of 50 bar both variables were statistically insignificant
(95 % confidence level, p<0.05) for extraction yield. The relationship between the
extraction yield, extraction temperature and static extraction time is depicted in Figure
5.3.1.
An increase in extraction temperature is reported to improve the efficiency of
extraction due to enhanced diffusion rate and solubility of analytes in solvents,
nevertheless, high extraction temperatures may simultaneously increase the degradation
rate of some interest compounds, such as anthocyanins, due to its high thermolability (Ju
and Howard, 2003), thus reducing the overall extraction yield.
Figure 5.3.1 - Three-dimensional response surfaces of the influence of extraction
temperature and static extraction time on the extraction yield.
70
5.3.2 Effects of process variables on the recovery of anthocyanins
The effects of extraction pressure, temperature and static extraction time on the
recovery of anthocyanins from jabuticaba skins were also evaluated. At a variable range of
50-100 bar, 40-80 ºC and 3-9 min, as occurred to the response variable extraction yield, the
extraction of anthocyanins was significantly (95 % confidence level, p<0.05) affected only
by the temperature. Figure 5.3.2.1a and 5.3.2.1b present the Pareto chart concerning the
effect (95% confidence level, p<0.05) of extraction variables on the recovery of
anthocyanins, where one can see that at 50-100 bar, 40-80 ºC and 3-9 min the extraction
temperature had a positive significant effect on this response variable, while at 80-120 ºC
and 9-15 min (at a fixed pressure of 50 bar) temperature and interaction between
temperature and static time affected negatively the extraction of anthocyanin pigments. The
relationship of the recovery of anthocyanins, extraction temperature and static extraction
time is depicted in Figure 5.3.2.2.
Arapitsas and Turner (2008) observed similar results for the extraction of
anthocyanins from red cabbage using also pressurized liquid extraction (PLE). Extraction
temperature and static extraction time ranges of 80-120 ºC and 6-11 min (at a fixed pressure
of 50 bar) were employed in their study; the optimal anthocyanin extraction was achieved
using short extraction time (7 min) and medium temperature (99 ºC). In agreement with our
results, according to the authors temperature seems to highly influence the anthocyanin
extraction. The temperature range employed (80-120 ºC) by Arapitsas and Turner was
decided after preliminary studies that had shown that higher temperatures (> 120 ºC)
caused anthocyanin degradation, and lower temperatures (< 80 ºC) gave poor extraction
efficiency of these pigments. In the present work, using a larger temperature range (40-120
ºC) it was observed analogous results (Figure 5.3.2.2). As the degradation rate of
71
anthocyanins is also time-dependent, high-temperature-short time extraction conditions
have been used successfully to obtain an anthocyanin-rich extract (Gizir et al., 2008).
Figure 5.3.2.1 - Effect (p<0.05) of extraction variables on the recovery of anthocyanins: a)
at 50-100 bar, 40-80 ºC and 3-9 min; b) at 80-120 ºC and 9-15 min at a fixed extraction
pressure of 50 bar (1, pressure; 2, temperature; 3, static time).
72
Figure 5.3.2.2 - Three-dimensional response surfaces of the influence of temperature and
static extraction time on recovery of anthocyanins.
Corroborating our results, Pompeu et al. (2009) have also concluded that
extraction temperature should not be increased indefinitely due to degradation of heat
sensitive phenolic compounds present in fruits of Euterpe oleracea as vegetable source,
such as anthocyanins. Thus, a maximum limit of temperature should be fixed and this will
depend on other factors (mainly time).
Usually, in the studies about the recovery of anthocyanins, a compromise
between high temperature and long time, that would increase the extraction, and low
temperature and short time, that would avoid the possibility of thermodegradation of
anthocyanins, is searched.
Based on the achievements of Petersson et al. (2010) studying the
extraction/degradation of anthocyanins from red onion during the pressurized liquid
73
extraction procedure we can elucidate what may have occurred in our study. Petersson and
co-authors verified that during all the extraction process, anthocyanin degradation and
extraction occur at the same time. Thus, it is suggested that in our work after a certain static
extraction time, a maximum level of anthocyanins will be reached, then degradation effects
would overcome the extraction effects, thus decreasing the anthocyanin level and
consequently the overall extraction yield.
5.3.3 Effects of process variables on the recovery of phenolic compounds
The effects of extraction pressure, temperature and static extraction time were
also studied for the extraction of other phenolic compounds. At 50-100 bar, 40-80 ºC and 39 min, extraction temperature exerted a significant influence again. Otherwise, extraction
pressure, static extraction time, their interaction, interaction between extraction pressure
and temperature and interaction between all three variables were as well statistically
significant. Figure 5.3.3.1a and 5.3.3.1b present the Pareto chart concerning the effect (95%
confidence level, p<0.05) of extraction variables on the recovery of total phenolic
compounds, where it is shown that at 50-100 bar, 40-80 ºC and 3-9 min the extraction
temperature, static extraction time, interaction between extraction pressure and static
extraction time and interaction between all three variables had a positive significant effect
on this response variable, while extraction temperature and interaction between extraction
temperature and pressure affected negatively the extraction of all phenolic compounds. At
80-120 ºC and 9-15 min (at a fixed extraction pressure of 50 bar) extraction temperature
and static extraction time continuously influenced positively the recovery of total phenolic
compounds, but their interaction affected negatively this response variable. The relationship
74
between the recovery of total phenolic compounds, extraction temperature and static
extraction time is depicted in Figure 5.3.3.2.
Figure 5.3.3.1 - Effect (p<0.05) of extraction variables on the recovery of total phenolic
compounds: a) at 50-100 bar, 40-80 ºC and 3-9 min; b) at 80-120 ºC and 9-15 min at a
fixed extraction pressure of 50 bar (1, pressure; 2, temperature; 3, static time).
75
Figure 5.3.3.2 - Three-dimensional response surfaces of the influence of temperature and
static extraction time on recovery of total phenolic compounds.
The results obtained in this work seem to be consistent with current scientific
literature. Herrero et al. (2010) extracting phenolic compounds from rosemary using PLE
and ethanol as solvent also verified that the highest the extraction temperature, the highest
the phenolic compounds extraction. Four temperatures (50, 100, 150 and 200 ºC) were
investigated at a constant extraction time and pressure. However, no significant difference
was observed between the total phenols extracted at 150 and 200 ºC, indicating that the
optimal extraction temperature is near 150 ºC. Indeed, Howard and Pandjaitan (2008) found
that near 150 ºC the extraction temperature should be set to effectively extract phenolic
compounds from Spinach by PLE method using ethanolic solvent (mixture of ethanol and
water; 70:30 v/v).
76
Differences in behavior between total phenolics and anthocyanins could be
explained by a higher susceptibility of the specific class of phenolic compounds
anthocyanins to high temperature (Cacace and Mazza, 2003). As in our work the main
objective is to find the PLE conditions that are more effective for extraction of thermolabile
anthocyanins, no higher temperature extraction level than 120 ºC were employed in order to
achieve the optimal extraction temperature for total phenolic compounds, which probably
could be as well near 150 ºC.
5.3.4 Optimization of the extraction process
The optimum PLE conditions for the extraction yields, anthocyanins and total
phenols from jabuticaba skins within the experimental variable ranges employed are
presented in the Table 5.3.4.1.
Table 5.3.4.1 - Optimum PLE conditions for the extraction yields, anthocyanins and total
phenols from jabuticaba skins
Optimum PLE conditions
Response Variables
Pressure (bar)
Temperature
Static
(ºC)
Extraction
Time (min)
Extraction yield (%)
Anthocyanins
(mg
Cy-3-glucoside/g
48-52
75-87
8-11
dry 48-51
78-82
8-10
118-120
14-15
material)
Total phenols (mg of GAEs/g dry material)
48-50
77
Considering the informations regarding the extraction yield and anthocyanin
extraction, the use PLE extraction pressure of 50 bar, temperature of 80 °C and static
extraction time of 9 min can result in optimal extraction yield (13.263 %) and anthocyanins
(2.139 mg Cy-3-glucoside/g dry material). On the other hand, at these conditions the total
phenols was 7.976 mg of gallic acid equivalents (GAEs)/g dry material; 2.34-fold lower
than the maximum total phenols (obtained at PLE extraction pressure of 50 bar,
temperature of 120 ºC and static extraction time of 15 min). The use of these PLE
conditions could be the most appropriate processing conditions to obtain a great amount of
extract with a high content of anthocyanins from jabuticaba skins.
It is well known that the performance of each technique in terms of anthocyanin
and other phenolic compounds extraction and yield produced could be effectively
compared minimizing the possible strong effects of the origin, year of production, preprocessing storage conditions and treatments of the sample (Al-Farsi et al., 2005;
Dourtoglou et al., 2006; Gizir et al., 2008). Thus, the extraction process at the selected
optimum PLE conditions was compared to conventional Low Pressure Liquid Extraction
(LPLE) using the same raw material and solvent. The experimental results demonstrated
that Pressurized Liquid Extraction procedure was much more effective in extracting
anthocyanins and other phenolic compounds from jabuticaba skins (Table 5.3.4.2). Similar
values of extraction yield were obtained by both extraction methods, however 2.15 and
1.66-fold more anthocyanins and phenolic compounds, respectively, were extracted using
PLE, compared to conventional LPLE. In both extraction processes the solid is stationary
and the solvent flows through the bed containing jabuticaba skins, hence for a better
comparison we have preferred this conventional LPLE. Even though, in the LPLE process
78
the solvent/solution is pumped continuously through the biomass to increase the efficiency
of intraparticle mass transfer of the phenolic compounds extraction was lower than when
using PLE process. This fact can be attributed to the use of combined pressure and
temperature during the PLE extraction process. Several authors have demonstrated that this
combination enhances the extraction of anthocyanins and other phenolic compounds from
different sources (Ju and Howard, 2003; Seabra et al., 2010; Corrales et al., 2009; Xu et al.,
2010).
Table 5.3.4.2 - Predicted and experimental values of responses under optimum PLE
conditions (50 bar, temperature of 80 °C and static extraction time of 9 min) and
experimental values responses obtained by Conventional Low Pressure Liquid Extraction
(LPLE)
PLE Values
Response Variables
Extraction yield (%)
LPLE Values
Experimental
Experimental
(mean ± SD)
(mean ± SD)
13.26
13.0 ± 0.9
12.1 ± 0.8
2.14
2.4 ± 0.5
1.2 ± 0.2
7.98
7.8 ± 0.4
4.8 ± 0.1
Predicted
Anthocyanins (mg Cy3-glucoside/g dry
material)
Total phenols (mg of
GAEs/g dry material)
According to our previous studies using the same extraction solvent and raw
material (fresh), the recovery of anthocyanins and other phenolic compounds using other
extraction methods, such as soxhlet, ultrasound-assisted and agitated solvent extraction,
79
were lower than of those obtained in this work (Santos et al., 2010). This might be due to
the adverse effects of drying (pre-processing treatment) of the sample on anthocyanins and
other phenolic compounds present in the jabuticaba skins. Gizir et al. (2008), comparing the
anthocyanin content of dried black carrot, verified that the anthocyanin content of dried
samples was lower than of fresh samples, indicating
that during the drying process
significant anthocyanin degradation occurred. Since this variation was identified, Gizir and
co-authors also focused on comparing their PLE extracts with extracts obtained by other
extraction methods using the same dried carrot sample. Herrero et al. (2010) have adopted
identical procedure in their studies as well due to the similar reasons.
In another previous study by our research group anthocyanins and other
phenolic compounds were extracted from fresh jabuticaba skins using the same LPLE
apparatus and solvent used in this work, although the results in terms of anthocyanin and
total phenolic compounds extraction were 88.15 and 58.84 % higher than those obtained
here, respectively (Santos et al., 2009), strengthening our hypothesis.
The promising use of pressurized ethanol for the extraction of anthocyanins and
phenolic compounds from jabuticaba skins gives the possibility to use a faster, more
efficient and more environmentally friendly technique. In fact, the optimized PLE
extraction was carried out in less than 21 min using small amounts of solvent (20 mL),
while the other low pressure conventional extraction methods took at least 2 h and used
2.25 times more ethanol in each run. By analysis of the experimental extraction kinetics
curves (Figure 5.3.4.1) it was possible to further reduce the PLE extraction time (reducing
the dynamic extraction time) and consequently reducing the amount of solvent used.
80
Figure 5.3.4.1 - Kinetics curves: a) overall extraction yield; b) recovery of anthocyanins; c)
recovery of total phenolic compounds under optimum PLE conditions (50 bar, temperature
of 80 °C and 9 min of static extraction time).
81
Figure 5.3.4.1 shows that at the beginning the anthocyanin and total phenolic
compounds extraction increases with increasing time, reflecting a faster solubility of
anthocyanins and other phenolic compounds into the unsaturated extraction solutions. After
a period of time the extraction was maximized into a steady-state extraction, indicating that
the mobility of anthocyanins and other phenolic compounds from jabuticaba skins into
extraction solution approaches zero in the remaining time. From an industrial point of view,
the operational time plays an important role also in the manufacturing cost estimative, once
it is related to the number of extraction cycles that can be performed by the extraction unit
(Rosa and Meireles, 2005).
5.4 Conclusions
The results presented on this contribution show the possibility to recover
phenolic compounds, such as anthocyanins, from jabuticaba skins using an environmentally
clean extraction technique. The results showed that extraction yield, anthocyanin and other
phenolic compounds extraction were most positively affected by extraction temperature.
Anthocyanin extraction was also affected by static extraction time indicating that higher
temperatures (> 80 ºC) and higher static extraction time (> 9min) caused anthocyanin
degradation, and lower temperatures and static extraction times gave poor extraction
efficiency of the pigments. Total phenolic compounds extraction was also affected by other
variables and their interactions, nevertheless optimum variable conditions were not
achieved using the variable ranges employed in this work. The use of PLE conditions set at
50 bar and 80 °C under a static extraction time of 9 min was selected for further economical
evaluation analysis since it produce the highest amount of extract with the highest content
82
of anthocyanins. Similar values of extraction yields were obtained by LPLE and PLE at
optimized conditions, however 2.15 and 1.66 more anthocyanins and total phenolic
compounds, respectively, were extracted using PLE. The experimental values agreed with
the predicted values, thus indicating suitability of the model employed and the success of
Response Surface Methodology (RSM) in optimizing the extraction conditions.
Acknowledgements
The authors are grateful to CNPq for financial support. Diego T. Santos thanks
CNPq (141894/2009-1) for the doctorate fellowship.
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Antioxidant Activity, Anthocyanins, Carotenoids, and Phenolics of Three Native Fresh and
Sun-Dried Date (Phoenix dactylifera L.) Varieties Grown in Oman. Journal of Agricultural
and Food Chemistry, 53 (19), 7592-7599.
Arapitsas, P., & Turner, C. (2008). Pressurized solvent extraction and monolithic columnHPLC/DAD analysis of anthocyanins in red cabbage. Talanta, 74, 1218-1223.
Cacace, J.E., & Mazza, G. (2003). Optimization of Extraction of Anthocyanins from Black
Currants with Aqueous Ethanol. Journal of food science, 68(1), 240-248.
Corrales, M., García, A.F., Butz, P., & Tauscher, B. (2009). Extraction of anthocyanins
from grape skins assisted by high hydrostatic pressure. Journal of Food Engineering, 90,
415-421.
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Dourtoglou, V.G., Mamalos A., & Makris, D.P. (2006). Storage of olives (Olea europaea)
under CO2 atmosphere: Effect on anthocyanins, phenolics, sensory attributes and in vitro
antioxidant properties. Food Chemistry, 99(2), 342-349.
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by UV–visible spectroscopy. In R. E. Wrolstad (Ed.) Current protocols in food analytical
chemistry. John Wiley & Sons, New York.
Gizir, A.M., Turker, N., & Artuvan, E. (2008). Pressurized acidified water extraction of
black carrot [Daucus carota ssp sativus var. atrorubens Alef.] anthocyanins. European
Food Research and Technology, 226, 363-370.
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of Chromatography A, 1217, 2512-2520.
Howard, L., & Pandjaitan, N. (2008). Pressurized Liquid Extraction of Flavonoids from
Spinach. Journal of food science, 73(3), C151-C157.
Ju, Z.Y., & Howard, L.R. (2003). Effects of solvent and temperature on pressurized liquid
extraction of anthocyanins and total phenolics from dried red grape skin. Journal of
Agricultural and Food Chemistry, 51, 5207-5213.
Petersson, E.V., Liu, J., Sjöberg, P.J.R., Danielsson, R., & Turner, C. (2010). Pressurized
Hot Water Extraction of anthocyanins from red onion: A study on extraction and
degradation rates. Analytica Chimica Acta, 663, 27-32.
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phenolic antioxidants from fruits of Euterpe oleracea using Response Surface
Methodology. Bioresource Technology, 100, 6076-6082.
Richter, B.E., Jones, B.A., Ezzell, J.L., & Porter, N.L. (1997). Accelerated solvent
extraction: a new technique for sample preparation. Analytical Chemistry, 68, 1033-1039.
Rosa, P.T.V., & Meireles, M.A.M. (2005). Rapid estimation of manufacturing cost of
extracts obtained by supercritical fluid extraction. Journal of Food Engineering, 67, 235240.
Santos, D.T., & Meireles, M.A.A. (2009). Jabuticaba as a source of functional pigments.
Pharmacognosy Reviews, 3 (5), 127–132.
Santos, D.T., Veggi, P.C, & Meireles, M.A.A.
(2010). Extraction of antioxidant
compounds from Jabuticaba (Myrciaria cauliflora) skins: Yield, composition and
economical evaluation. Journal of Food Engineering, 101, 23-31.
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85
86
CAPÍTULO 6 - OPTIMIZATION OF BIOACTIVE COMPOUNDS EXTRACTION
FROM JABUTICABA (MYRCIARIA CAULIFLORA) SKINS ASSISTED BY HIGH
PRESSURE CO2
Diego T. Santos and M. Angela A. Meireles
Trabalho submetido ao periódico Innovative Food Science and Emerging Technologies.
87
Key words
Anthocyanins, Phenolic compounds, Jabuticaba skins, Myrciaria cauliflora, Optimization,
High Pressure Carbon Dioxide Assisted Extraction
Abstract
Important bioactive compounds from Brazilian jabuticaba skins were effectively extracted
by High Pressure Carbon Dioxide Assisted Extraction (HPCDAE). Statistical design was
used to optimize the extraction variables: extraction pressure (65-135 bar), temperature (4080 ºC) and volume ratio of solid–liquid mixture/pressurized CO2 ( RS − L / CO2 (%) (20-80 %).
The analysis performed to predict the optimum values for the extraction variables, in order
to obtain the conditions that results in an extract with high anthocyanin (2.2 ± 0.3 mg
cyanidin-3-glucoside/g dry skins) and phenolic compounds contents (13 ± 1 mg gallic acid
equivalents/g dry skins), gave as best conditions 117 bar extraction pressure, 80 ºC
extraction temperature and 20% volume ratio of solid–liquid mixture/pressurized CO2
( R S−L / CO 2 (%) ). Compared to Pressurized Liquid Extraction (PLE) and to control
experiment the results obtained using optimum HPCD Assisted-Extraction conditions was
much more effective and faster in extracting total anthocyanins and phenolic compounds.
Industrial relevance: Industrially, there is an increasing demand for faster extraction
procedures with reduced organic solvent consumption to lower pollution burden. HPCD
Assisted-Extraction combines the advantages of enhanced mass transfer rates increasing
secondary metabolite diffusion from the vegetable matrix into the environmentally friendly
solvent extraction. High pressure extraction methods, such as HPCD Assisted-Extraction,
has other advantages that should be considered, such as the fact that native enzymes, which
degrade phenolic compounds, are inhibited by extraction pressure increase and CO2
addition, and that processed vegetable materials do not require additional sterilization steps.
88
6.1 Introduction
In recent years, the desire for a healthier diet allied with the increasing concern
of consumers over the use of synthetic additives in food has pushed the food industry to
search for new sources of natural pigments (Montes, Vicário, Raymundo, Fett, & Heredia,
2005). In Brazil a potential source of natural pigments is jabuticaba skins. Jabuticaba
(Myrciaria cauliflora) is grape-like in appearance and texture, although its skin is thicker
and tougher. This fruit has a dark purple to almost black skin color due to a high content of
anthocyanins that cover a white gelatinous flesh inside (Santos, Veggi, & Meireles, 2010).
Anthocyanins, coloured natural compounds can be considered potential
replacements for the synthetic food dyes. Moreover, the proved biological activities of
anthocyanins, related to the prevention of a number of degenerative diseases (Santos &
Meireles, 2009), provide added benefits to food products dyed with these compounds.
Anthocyanins are traditionally extracted from plant materials by conventional
methods, such as solid–liquid extraction, using sometimes large amounts of organic
solvent, for example methanol, ethanol and acetone or their aqueous solutions containing a
small amount of acid to maintain a low pH, where anthocyanins are in their most stable
flavylium form (Gizir, Turker, & Artuvan, 2008). Nevertheless, there is an increasing
demand for faster extraction procedures with reduced organic solvent consumption and
lower pollution burden (Wang & Weller, 2006).
Anthocyanins are particularly unstable, being especially sensitive to light,
alkaline pH, and heat (Stintzing, Stintzing, Carle, Frei, & Wrolstad, 2002). Therefore, to
avoid degradation during anthocyanin extraction, high-temperature-short time extraction
conditions have been successfully employed (Arapitsas & Turner, 2008).
Thus, it is a key focus to develop novel extraction methods with faster
extraction rates and higher anthocyanin extraction yields. An efficient extraction should
maximize anthocyanin recovery with minimal degradation using environmentally friendly
technologies. In this way, the use of Pressurized Liquid Extraction (PLE®), sometimes
referred to as Pressurized Solvent Extraction (PSE®) and Accelerated Solvent Extraction
89
(ASE®) using acidified water or an acidified alcohol and their mixtures (Petersson, Liu,
Sjöberg, Danielsson, & Turner, 2010; Ju & Howard, 2003), Supercritical Fluid Extraction
(SFE) using carbon dioxide in combination with water and/or an alcohol as co-solvent
(Seabra, Braga, Batista, & Sousa, 2010; Ghafoor, Park, & Choi, 2010), High Hydrostatic
Pressure Assisted-Extraction (HHPAE), Pulsed Electric Fields Assisted-Extraction
(PEFAE) using water/ethanol mixture (Corrales, Toepfl, Butz, Knorr, & Tauscher, 2008),
High Pressure Carbon Dioxide Assisted-Extraction (HPCDAE) using acidified water (Xu et
al., 2010), among others, is an attractive alternative.
In this study, two different emerging novel technologies, HPCDAE and PLE,
were used for the extraction of anthocyanin-based pigment and other phenolic compounds
from jabuticaba skins. Xu et al. (2010) have demonstrated for the first time that the
explosive effect of high pressure CO2 (HPCD), besides inactivate microorganisms and
enzymes, can also strengthen anthocyanin extraction process by its superior abilities in cell
membrane modification, intracellular pH decrease, disordering of the intracellular
electrolyte balance and removal of vital constituents from cells and cell membranes. As the
quality of an extract is related to its chemical composition, and then, to its functional
properties, an efficient extraction method should maximize the extraction of anthocyanins
and other phenolic compounds synchronously (Santos, Veggi, & Meireles, 2010).
Therefore, the aim of this work was to optimize HPCDAE variables such as
extraction pressure, temperature and volume ratio of solid–liquid mixture/pressurized CO2
( RS − L / CO2 (%)) for the maximum recovery of anthocyanins and total phenolic compounds by
acidified water. HPCDAE process was optimized by using Response Surface Methodology
(RSM) and compared to PLE at the optimum extraction pressure and temperature
conditions.
90
6.2 Material and methods
6.2.1 Plant material
Jabuticaba fruits (Myrciaria cauliflora), harvested from a plantation in the State
of São Paulo, Brazil, were acquired from a fruit and vegetable market centre (CEASACampinas, Brazil). Immediately after acquiring, the fruits were manually peeled and the
jabuticaba skins were dried for a few hours at 45 ºC in an oven with forced air circulation
(Marconi, MA 035/1, Piracicaba, São Paulo, Brazil). The dried skins (59.17 % moisture)
were stored in the dark in a domestic freezer (-10 ºC) (Double Action, Metalfrio, São Paulo,
Brazil) until extraction.
6.2.2 High Pressure Carbon Dioxide Assisted-Extraction (HPCDAE) system
The diagram of the HPCDAE system is shown in Figure 6.2.1. The HPCDAE
system was designed and assembled at LASEFI/DEA/FEA (School of Food
Engineering)/UNICAMP (University of Campinas). All connections within the system
were made of stainless steel tubes (1/16” and 1/8”). The stainless steel pressure vessel with
a volume of 500 mL (6.8 cm internal diameter) was designed to withstand a pressure of 200
bar and temperature of 250 ºC. The vessel temperature was maintained by a heating water
bath (Marconi, MA 127BO, Piracicaba, Brazil). One thermocouple with a digital
temperature display (Autonics, T4WM, Bucheon, South Korea) was fixed in the vessel lid
to monitor the CO2 temperature in the upper part of the vessel. A Bourbon type manometer
(250 ± 1 bar, Record, Classe:A2, São Paulo, Brazil) was connected by 1/8” tubes to the
vessel to monitor the vessel pressure. Pressure was controlled by a back pressure regulator
(Tescom, model # 26-1761-24-161, ELK River, USA). Before reaching the air driven liquid
pump (Maximator Gmbh, PP 111, Zorge, Germany), the CO2 was cooled under -10 °C by a
thermostatic bath (Marconi, MA-184, Piracicaba, Brazil). The system also includes a
micrometric valve with a heating system with a digital temperature controller (Novus,
91
N480D, Porto Alegre, Brazil) to avoid Joule-Thomson freezing effect during rapid
depressurization that can lead to clogging of the throttling device; a CO2 glass float
rotameter (0.15-2.2 kg/h of CO2 at 1.013bar/20 ºC, ABB, 16/286A/2, Warminster, USA); a
flow totalizer (LAO, model G0,6, Osasco, Brazil); an online filter (0.5 μm, Swagelok,
Solon, USA); a safety valve (400 bar, Swagelok, R3A-G, Solon, USA); 2 Bourbon type
manometers of 500 ± 3 bar and 1 of 100 ± 0.5 bar (Record, Classe:A2, São Paulo, Brazil).
3
4
2
7
3
3
4
11
5
4
4
6
1
4
12
3
10
11
9
8
Figure 6.2.1 - Diagram of High Pressure Carbon Dioxide Assisted-Extraction (HPCDAE)
system. 1 CO2 cylinder; 2 CO2 filter; 3 Manometers; 4 Valves; 5 Thermostatic bath; 6
Pump; 7 Back pressure regulator; 8 Heating bath; 9 High pressure vessel; 10
Thermocouple; 11 Temperature controllers; 12 Micrometric valve with a heating system.
6.2.3 Extraction Procedures
6.2.3.1 High Pressure Carbon Dioxide Assisted-Extraction (HPCDAE)
92
First, the pressure vessel was heated to the required temperature (40-80 ºC). A
volume (90-360 cm3) of acidified distilled water [pH 2.5 ± 0.2 by hydrochloric acid (Synth,
Diadema, Brazil)] preheated using a heating plate (Fisaton, model 752A, São Paulo, Brazil)
was filled into the vessel. Afterwards a given weight of jabuticaba skins (9-36 g) was
placed into the vessel and the cover of the vessel was tightened. The vessel was pressurized
with CO2 (99.9 %, Gama Gases Especiais Ltda., Campinas, Brazil) by the pump to the
required pressure (65-135 bar), which was held for the determined time (5, 10, 20, 30 or 45
min). Then the depressurization was performed by releasing CO2 into the atmosphere using
the pressure relief micrometric valve with a heating system (80 ºC). After completion of
HPCD Assisted-Extraction, a sample (10 cm3) of the aqueous extract was collected into a
sample bottle immersed in ice water (5 °C) to prevent anthocyanin and phenolic
compounds degradation. All the aqueous extracts were stored (- 10 ºC) in the dark until
analysis. The vessel covering and uncovering time were 7 and 4 min, respectively; the
pressurization and depressurization time were in the range of 1–4 and 7-8 min, respectively,
depending of the pressure employed.
The value change of the volume of solid–liquid mixture (S-L) at the
experimental conditions evaluated was neglected since the density of water varied slightly
(from 0.97464 to 0.99801 g/mL) (http://webbook.nist.gov/chemistry/fluid).
To determine the effects of extraction pressure, temperature and volume ratio of
solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) on the recovery of anthocyanins and
phenolic compounds, twenty extractions in random order were carried out. All the
extractions were performed in duplicate.
An extraction under atmospheric pressure as a control was performed at 80 oC
using the HPCDAE system without pressurizing it. A sample (1 mL) was taken out at 5, 10,
20, 30 or 45 min of extraction. No calculations were made to compensate for volume loss
93
due to sample collection, since the volume change of the extraction solvent had no effect on
the concentration of the extract.
For both HPCDAE and control experiment, the ratio of solid vs. liquid ratio
was 1:10 (1 g of jabuticaba skins:10 mL acidified water).
6.2.3.2 Pressurized Liquid Extraction (PLE)
Four and one half grams of jabuticaba skin pieces were placed in the 6.57 mL
extraction cell (Thar Designs, Pittsburg, USA) containing a syntherized metal filter at the
cell inlet and outlet. The cell containing the sample was heated by an electrical heating
jacket at a desired temperature, filled with extraction solvent [acidified distilled water pH
2.5 ± 0.2 by hydrochloric acid (Synth, Diadema, Brazil)] and then pressurized. The
extraction solvent was pumped by a HPLC pump (Thermoseparation Products, Model
ConstaMetric 3200 P/F, San Jose, USA) into the extraction cell until the required pressure
was obtained. All connections within the system were made of stainless steel tubes (1/16”
and 1/8”).
During 5 min the plant material was placed in the heating system to ensure that
the extraction cell would be at the desired temperature (80 ºC) at the filling and
pressurization steps. After pressurization, the jabuticaba skins with pressurized solvent
were kept statically at the desired pressure (117 bar) for a desired time (5 min). The static
extraction time of 5 min was decided after preliminary studies that had shown that higher
times caused anthocyanin degradation, and lower times (< 5 min) gave poor extraction
efficiency of the desired compounds (anthocyanins and other phenolic compounds).
Thereafter, carefully the blocking and micrometric valve were opened keeping the pressure
constant to the desired flow rate (1 mL/min); the extraction cell was rinsed with fresh
extraction solvent during 45 min. Samples were collected at 5, 10, 15, 20, 30 and 45 min in
94
glass flasks. After PLE, aqueous extracts were rapidly cooled to 5 °C in ice water to prevent
anthocyanin and phenolic compounds degradation. All aqueous extracts were stored (- 10
ºC) in the dark until analysis.
6.2.4 Extract Characterization
6.2.4.1 Total Monomeric Anthocyanins (TMA) content
The Total Monomeric Anthocyanin (TMA) content was determined using the
pH differential method described by Giusti & Wrolstad (2001), which relies on the
structural transformation of the anthocyanin chromophore as a function of pH. A UV–Vis
spectrophotometer (Hitachi, model U-3010, Tokyo, Japan) was used for spectral
measurements at maximum absorbance wavelength (approximately 512 nm) and 700 nm,
using distilled water as blank. Two dilutions of the same sample were prepared: one with
hydrochloric acid/potassium chloride buffer pH = 1.0 and the other with sodium
acetate/acetic acid buffer pH = 4.5. The pH values of the buffers were measured using a
pH-meter (Digimed, model DM-22, São Paulo, Brazil) calibrated with buffers at pH 4.01
and 6.86 and they were adjusted with HCl (99.5 % Ecibra, Santo Amaro, Brazil). Aliquots
of aqueous extract were brought to pH 1.0 and 4.5; 15 min later, the absorbance of each
equilibrated solution was measured at the maximum absorption wavelength and 700 nm for
haze correction using a 1 cm path length glass cells (l). The dilution factor (DF) was
determined (final volume per original sample volume). The difference in absorbance values
at pH 1.0 and 4.5 is directly proportional to the TMA concentration. The anthocyanin
content was calculated as cyanidin-3-glucoside (MW = 449.2 g/mol and ε = 26.900
L/mol.cm) and the results were expressed as mg Cy-3-glucoside/g dry material. The
95
absorbance of the diluted sample (A) and the TMA were calculated with Equations (1) and
(2). The anthocyanin content (mg Cyanidin-3-glucoside/g dry weight) was calculated with
the Equation (3), where V (L) is the final volume of the aqueous extracts, m (g) is the
weight of jabuticaba skins (0 % moisture).
A = ( A max − A 700 ) pH 1, 0 − ( A max − A 700 ) pH 4,5
TMA ( mg / L ) =
A x MW x DF
X 1000
εxl
(1)
(2)
Anthocyanin content (mg Cyanidin − 3 − glu cos ide / g dry weight ) =
TMA x V
m
(3)
6.2.4.2 Total phenolic content
Total phenolic content was estimated using the Folin-Ciocalteau method for
total phenolics, based on a colorimetric oxidation/reduction reaction of phenols (Singleton
& Rossi, 1965). Briefly, 1 cm3 of sample (the sample preparation was done by diluting the
aqueous extract in distillated water) was mixed with 1 cm3 of Folin and Ciocalteu’s phenol
reagent. After 3 min, 1 cm3 of saturated sodium carbonate solution (50 % w/w) was added
to the mixture and the volume adjusted to 10 cm3 with distilled water. The reaction was
kept in the dark for 90 min at room temperature, after which the absorbance was read at 725
nm with a UV–Vis Spectrophotometer Hitachi, model U-3010 (Tokyo, Japan). For control
sample, 1 cm3 of distilled water was taken. The results were calculated on the basis of the
calibration curve of gallic acid (GAE, milligrams of gallic acid equivalents/mL) and
expressed as milligrams of gallic acid equivalents/g dry material (Equation 4), where V
(mL) is the final volume of the aqueous extracts, m (g) is the weight of jabuticaba skins (0
% moisture), DF is the dilution factor (final volume per original sample volume).
Phenolic content (mg gallic acid equivalents / g dry weight ) =
96
GAE (mg / mL) x DF x V
m
(4)
6.2.5 Statistical analysis
All the experimental results obtained were expressed as means ± SD. Statistical
analyses were performed using analyses of variance (ANOVA) with consequent
experimental design. The mean values were considered significantly different when p<0.05.
‘Statistica’ software (release 7, StatSoft, Tulsa, USA) was used to calculate the effects of
the extraction pressure, temperature and volume ratio of solid–liquid mixture/pressurized
CO2 ( R S−L / CO 2 (%) ) by a 23 full factorial design on the recovery of anthocyanins and
phenolic compounds employing extraction pressure, temperature and volume ratio of solid–
liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) ranges of 65-135 bar, 40-80 ºC and 20-80
%, respectively. The 23 full factorial design to optimize the HPCD Assisted-Extraction of
total anthocyanins and phenolic compounds from jabuticaba skins is represented in Table
6.3.1.1. The optimum extraction conditions were estimated through regression analysis and
three dimensional response surface plots of the independent variables and each dependent
variable. Response surface analysis was also applied on the data from 23 full factorial
design for modeling and prediction of optimum conditions of HPCD Assisted-Extraction
for total anthocyanins and total phenols from jabuticaba skins.
6.2.6 Determination of experimental extraction kinetics curves and parameters
The experimental extraction kinetics curves plotting and the determination of
experimental values of the steady-state extraction Y*, the time t* to reach the Y* and the
M* (the mass transfer rate) for the recovery of anthocyanins and phenolic compounds were
performed with Office 2003 Excel (Microsoft Co., Redmond, USA).
6.3 Results and discussion
6.3.1 Effects of process variables on recovery of anthocyanins
The experimental values of anthocyanin content of the extracts at various
experimental conditions are presented in Table 6.3.1.1. Regression analysis was performed
on the experimental data and the coefficients of model were evaluated for significance. The
97
regression analysis of the data showed that the recovery of anthocyanins was significantly
(p<0.05) affected by the extraction pressure, temperature and their interactions, while the
effect of volume ratio of solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) was not
significant. The relationship of the anthocyanin content, extraction pressure and
temperature is depicted in Figure 6.3.1.1 and it was linear with R2 value of 0.981. An
increase in either pressure or temperature, while the second variable remains constant,
results in enhancement of the anthocyanin extraction (Figure 6.3.1.2). Nevertheless, the
interaction between them had a negative effect on the recovery of anthocyanins. The
synchronous increase of the extraction pressure and temperature possibly might enhance
the degradation of the anthocyanins extracted. Combined temperature/pressure treatments
have enhanced the anthocyanin degradation and formation of condensation products in
wines which contribute to colour, organoleptic and nutritional changes (loss of antioxidant
capacity) (Corrales, Butz, & Tauscher, 2008). The relationship between process variables
and the anthocyanin content (mg cyaniding-3-glucoside/g dry material) is presented in
Equation (5) by omitting all regression coefficients that were insignificant.
Y1 = − 3.46311 + 0.03743.X 1 + 0.07766.X 2 − 0.00049.X 1 .X 2
(5)
In Equation 5, Y1 is the anthocyanin content (mg cyaniding-3-glucoside/g dry
material) in jabuticaba skins extract, X1 is the extraction pressure (bar), X2 is the extraction
temperature (º C). The equation was based on the data of regression coefficients presented
in Table 6.3.1.2.
A positive effect of the high extraction pressure and temperature, combined
or not, on the recovery of anthocyanins using other kinds of static (batch) extraction was
also observed. Corrales, Toepfl, Butz, Knorr, & Tauscher (2008) using a mixture of water
and ethanol (50:50, v/v) as extraction solvent under High Hydrostatic Pressure (HHP)
(6000 bar) at 70 °C have proposed that HHP combined with temperature possibly leaded to
a decrease in the dielectric constant of water and in the pH of the solvent during the
extraction, enhancing the extraction of anthocyanins from grape by-products. Using a
98
closed reactor vessel under agitation using a mixture of acidified water and ethanol (95:5
v/v) at 110 °C as extraction solvent for the anthocyanin recovery from red onion, Petersson,
Liu, Sjöberg, Danielsson, & Turner (2010) have observed, even though extraction and
degradation effects compete during the extraction process, that high temperature can exert a
higher influence on the extraction rate than on the degradation rate, improving anthocyanin
extraction.
Table 6.3.1.1 - 23 full factorial design for HPCD Assisted-Extraction from jabuticaba skins
and the total anthocyanin and phenolic contents of the extracts
HPCD Assisted-Extraction conditions
Experiment Pressure Temperature
R S−L / CO 2 (%)
Analytical results
Anthocyanin
Phenolic
content
content
(bar)
(ºC)
1
65
40
20
0.8 ± 0.1
3.5 ± 0.2
2
135
40
20
2.04 ± 0.08
3.94 ± 0.07
3
65
80
20
2.55 ± 0.01
14.0 ± 0.7
4
135
80
20
2.6 ± 0.7
14.4 ± 0.3
5
65
40
80
0.6 ± 0.1
2.6 ± 0.3
6
135
40
80
1.81 ± 0.01
4.81 ± 0.01
7
65
80
80
2.1 ± 0.1
12.3 ± 0.9
8
135
80
80
2.5 ± 0.3
12.4 ± 0.2
9
100
60
50
1.33 ± 0.01
4.6 ± 0.7
10
100
60
50
1.34 ± 0.01
4.3 ± 0.6
99
Figure 6.3.1.1 - Three-dimensional response surfaces of the influence of extraction pressure
and temperature on recovery of anthocyanins.
Figure 6.3.1.2 - Two-dimensional response surfaces of the influence of extraction pressure
and temperature on recovery of anthocyanins.
100
Table 6.3.1.2 - Regression coefficients of the model for the response variables
Factor
Regression coefficients
Anthocyanin content
Phenolic content
Mean
-3.46311
-8.51056
(1) Pressure (bar)
0.03743
0.00253
(2) Temperature (º C)
0.07766
0.27832
(3) R S−L / CO 2 (%)
0.00682
-0.04071
1 by 2
-0.00049
0.00005
1 by 3
-0.00010
0.00077
2 by 3
-0.00025
0.00022
1*2*3
0.00000
-0.00001
The pH decrease that occurs due to the generation of “in situ” carbonic acid
and/or alkyl carbonic acid, when the CO2, which is a relatively soluble gas in water
(Enomoto, Nakamura, Nagai, Hashimoto, & Hakoda, 1997), is inserted into the HPCDAssisted Extraction system modifying the extracting solvent (acidified water), can have had
a positive impact on the anthocyanin extraction and stability from jabuticaba skins, besides
increasing cell membranes permeability, which leads to higher diffusivities (Türker &
Erdogdu, 2006). According to Xu et al. (2010), in HPCD-Assisted Extraction process, there
are five forms associated with CO2, including supercritical CO2, H2CO3 and its dissociated
products, H+, HCO3-3, and CO3-2, which possibly play different roles in the extraction of
anthocyanins, until now not fully understood.
6.3.2 Effects of process variables on recovery of phenolic compounds
The analytical results of phenolic content in jabuticaba skins extract are shown
in Table 6.3.1.1. The effect of extraction temperature was highly significant (p<0.001) on
the extraction of phenolic compounds, indicating that the main extraction variable for
101
phenolic compounds from this vegetable source is extraction temperature. The effects of
extraction pressure, volume ratio of solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) )
and the interaction between extraction temperature and volume ratio of solid–liquid
mixture/pressurized CO2 ( R S−L / CO 2 (%) ) were also significant (p<0.05). The relationship
between phenolic content (mg gallic acid equivalents/g dry material), extraction pressure
and temperature is depicted in Figure 6.3.2.1 and it was linear with R2 value of 0.996. As
occurred to anthocyanin recovery, an increase in either the pressure or the temperature,
while the second variable remains constant, results in enhancement of the phenolic
compounds extraction (Figure 6.3.2.2). The interaction between extraction temperature and
volume ratio of solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) had a negative effect
on the recovery of phenolic compounds. Equation (6) shows the relationship between
process variables for the extraction of total phenolic compounds by omitting all regression
coefficients that were insignificant.
Y2 = − 8.51056 + 0.00253.X1 + 0.27832.X 2 − 0.04071.X 3 + 0.00022.X 2 .X 3
(6)
In Equation 6, Y2 is the phenolic content (mg gallic acid equivalents/g dry
material) in jabuticaba skins extract, X1 is the extraction pressure (bar), X2 is the extraction
temperature (º C), X3 is the volume ratio of solid–liquid mixture/pressurized CO2
( R S−L / CO 2 (%) ). Equation (6) is based on the regression coefficients presented in Table
6.3.1.2.
102
Figure 6.3.2.1 - Three-dimensional response surfaces of the influence of extraction pressure
and temperature on the recovery of phenolic compounds.
Xu et al. (2010), using HPCD Assisted-Extraction and acidified water (pH 2.0 ±
0.2) for anthocyanin extraction from red cabbage observed similar results to those obtained
in the present study. For the extraction of this specific phenolic compound higher
temperature (40-60 °C) also increased its recovery. Moreover, it was observed that
increasing the volume of pressurized CO2 (decreasing the volume ratio of solid–liquid
mixture/pressurized CO2 ( R S−L / CO 2 (%) ) (16.5-81.2 %) also benefited the extraction
process. Xu et al. (2010) reported that higher volume of pressurized CO2 produced stronger
explosive effect during the fast decompression step, which probably can cause faster and
more effective mass transfer of the solutes from the plant matrix to the extracting solvent
by plant cell destruction in these processes (Enomoto, Nakamura, Nagai, Hashimoto, &
Hakoda, 1997). No different pressures were evaluated in their study, however comparing
103
the extraction under 100 bar to the extraction under atmospheric pressure (control
experiment) at the same extraction temperature, the results indicate that raising pressure
increased the extraction of phenolic compounds, corroborating our results.
Figure 6.3.2.2 - Two-dimensional response surfaces of the influence of extraction pressure
and temperature on the recovery of phenolic compounds.
High pressure extraction methods, such as HPCD Assisted-Extraction, have
other advantages that should be considered, such as the fact that native enzymes, which
degrade phenolic compounds, are inhibited by extraction pressure increasing and CO2
addition, and that processed vegetable materials do not require additional sterilization steps.
Furthermore, the absence of oxygen in the extraction cell is other advantage of this process,
because its presence can lead to structural changes in phenolic compounds that can, in turn,
result in altered properties (Seabra, Braga, Batista, & Sousa, 2010).
104
6.3.3 Optimization of the extraction process
As demonstrated before, extraction pressure, temperature and volume ratio of
solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) have different effects on each
response variable evaluated. As the quality of an extract is related to its chemical
composition, and then, to its functional properties, an efficient extraction method should
maximize the extraction of phenolic compounds with acceptable degradation of
anthocyanin pigments (Santos, Veggi, & Meireles, 2010).
The prediction of one set of optimal conditions for both response variables was
done by using desirability function approach. Figure 6.3.3.1 shows that all variables
(extraction
pressure,
extraction
temperature
and
volume
ratio
of
solid–liquid
mixture/pressurized CO2 ( R S−L / CO 2 (%) ) seem to influence the desirability to obtain the
desired extract (high anthocyanin and phenolic contents). In particular, these graphs show
that high desirability, within the experimental design values, has been achieved under
medium
pressure,
high
temperature
and
low
volume
ratio
of
solid–liquid
mixture/pressurized CO2 ( R S−L / CO 2 (%) ).
The analysis performed to predict the optimum values for the extraction
variables, in order to obtain the conditions that result in an extract with simultaneously high
anthocyanin and phenolic contents, gave as best conditions, 117 bar extraction pressure, 80
ºC extraction temperature and 20% volume ratio of solid–liquid mixture/pressurized CO2
( R S−L / CO 2 (%) ) (Figure 6.3.3.2).
HPCD Assisted-Extraction pressures in the range of 115-120 bar, extraction
temperatures of 79-80 °C, volume ratio of solid–liquid mixture/pressurized CO2
( R S−L / CO 2 (%) ) of 20 %, can result in optimal total anthocyanins (2.543 mg of Cy-3glucoside/g dry material) and total phenolic compounds (13.589 mg of gallic acid
equivalents/g dry material) from jabuticaba skins. The predicted results matched well with
the experimental results obtained using optimum extraction conditions which validated the
response surface methodology model with a good correlation (Table 6.3.3.1).
105
Figure 6.3.3.1 - Three-dimensional response surfaces of the influence of the extraction
variables on the desirability.
Figure 6.3.3.2 - Profiles of the predicted values and desirability of the extraction variables.
106
The positive characteristics of using HPCD for anthocyanin and phenolic
compounds extraction synchronously seem irrefutable when the results obtained using this
novel technique are compared to that obtained using others extraction methods and to
literature data. Compared to Pressurized Liquid Extraction (PLE) with the same solvent at
the same extraction pressure, temperature and time and to control experiment (using
HPCDAE system at the same extraction temperature and time; without pressurization) the
experimental results obtained using optimum HPCD Assisted-Extraction conditions was
much more effective in extracting anthocyanins and phenolic compounds from jabuticaba
skins (Table 6.3.3.1). Montes, Vicário, Raymundo, Fett, & Heredia (2005), aiming to
optimize the conventional solvent extraction conditions (24 h at 4 ºC) for the maximum
recovery of anthocyanins from jabuticaba skins, using different solvents acidified with
different types of acids at different pHs, obtained a maximum extraction of anthocyanins of
92.920 mg/kg wet material, while our study found at the optimum HPCD AssistedExtraction conditions an experimental value 9.83-fold higher (913.529 mg/kg wet material,
i.e. 2.237 mg/g dry material).
Cacace & Mazza (2002) have suggested that the reduction of the dielectric
constant of water into the range of intermediate-behavior solvents such as methanol or
ethanol, good solvents for the extraction of anthocyanins and other phenolic compounds,
can be achieved not only by modifying pressure and temperature, but also with the addition
of ions in acidified water. A lower dielectric constant reduces the energy required to
separate the solvent molecules and allows the solute molecules to enter between them
(Mackay & Mackay, 1981). Based on these research results it is suggested that the addition
107
of CO2 into the water during the HPCD Assisted-Extraction may improve the extraction by
modifying the solvent, thus resulting in increased solubility of the target compounds.
Table 6.3.3.1 - Predicted and experimental values of response variables under optimum
HPCDAE conditions (at 117 bar, temperature of 80 °C,
of 20 % and 20 minutes of
extraction) and experimental values of response variables obtained by control HPCDAE
experiment (at atmospheric pressure, temperature of 80 ºC and 20 minutes of extraction)
and by PLE experiment (at 117 bar, temperature of 80 ºC, 20 minutes of extraction and
flow rate of 1 mL of acidified water/min)
Response Variables
HPCDAE Values
HPCDAE
(control) Values
PLE Values
Predicted
Experimental
Experimental
Experimental
2.543
2.24 ± 0.3
1.6 ± 0.1
0.58 ± 0.02
13.589
12.90 ± 1.46
8.3 ± 0.6
3.6 ± 0.1
Recovery of
anthocyanins (mg of
Cy-3-Glucoside/g
dry material)
Recovery of phenolic
compounds (mg of
GAEs/g dry material)
6.3.4 Experimental extraction kinetics curves using optimum conditions
Figure 6.3.4.1 and 6.3.4.2, respectively, confirm that HPCD effectively
increases the extraction of anthocyanins and phenolic compounds. In general, the
experimental extraction kinetics curves for all extraction methods were characterized with
two distinct phases. In the first phase, at the beginning the bioactive compounds extraction
108
increases with increasing extraction time, reflecting a faster solubility of anthocyanins and
phenolic compounds into the unsaturated extraction solutions. In the second phase, the
extraction was maximized into the steady-state extraction Y*, indicating that the mobility
of anthocyanins and phenolic compounds from jabuticaba skins into extraction solution
approaches zero ((starting at t*) in the remaining time. An exception is observed to the
extraction kinetics curve of phenolic compounds by PLE (Figure 6.3.4.2); in the extraction
time range evaluated the steady-state extraction Y* is not reached.
4
mg Cyanidin-3-glucoside /
g dry material
3.5
3
2.5
2
1.5
1
0.5
0
0
5
10
HPCD Assisted-Extraction
15
20
25
Extraction time (min)
30
35
HPCD Assisted-Extraction (Control)
40
45
PLE
Figure 6.3.4.1 - Kinetics curves for the recovery of anthocyanins a) under optimum
HPCDAE conditions (117 bar, temperature of 80 °C and of 20 %; b) obtained by control
HPCDAE experiment (at atmospheric pressure, temperature of 80 ºC and 20 minutes of
extraction) and c) PLE experiment (at 117 bar, temperature of 80 ºC, 20 minutes of
extraction and flow rate of 1 mL/min of acidified water/min).
109
mg gallic acid equivalents /
g dry material
25
20
15
10
5
0
0
5
10
HPCD Assisted-Extraction
15
20
25
Extraction time (min)
30
35
HPCD Assisted-Extraction (Control)
40
45
PLE
Figure 6.3.4.2 - Kinetics curves for the recovery of phenolic compounds a) under optimum
HPCDAE conditions (117 bar, temperature of 80 °C and of 20 %; b) obtained by control
HPCDAE experiment (at atmospheric pressure, temperature of 80 ºC and 20 minutes of
extraction) and c) PLE experiment (at 117 bar, temperature of 80 ºC, 20 minutes of
extraction and flow rate of 1 mL/min of acidified water/min).
It is also observed that HPCD can accelerate the extraction process of
anthocyanins and phenolic compounds. The amount of anthocyanin and phenolic
compounds extracted during the same extraction time by HPCD Assisted-Extraction under
optimum conditions is higher than that of the PLE and extraction under atmospheric
pressure (control HPCDAE experiment) in the first phase, as well as the Y* of the HPCD
Assisted-Extraction in the second phase (Table 6.3.4.1). The mass transfer rates (M*) in the
first phase of the extraction kinetics curves for the recovery of anthocyanins and phenolic
compounds using HPCDAE were, respectively, 1.52- and 2.13- fold higher than those
110
obtained by HPCDAE (control) and 4.70- and not determinated- fold higher than those
obtained by PLE (Table 6.3.4.1).
Table 6.3.4.1 - Experimental values of the steady-state extraction Y*, the time t* to reach
the Y* and the mass transfer rate M* for the recovery of anthocyanins and phenolic
compounds
Extraction method
HPCDAE under optimum conditions
(117 bar,
For the
Y* (mg/g
t*
M* (mg/g
recovery of
dry material)
(min)
dry material.s)
anthocyanins
3.360
37
5.448
20.660
33
37.564
1.973
33
3.587
10.869
37
17.625
0.753
39
1.158
*n.d
*n.d
*n.d
temperature of 80 °C and R S−L / CO 2 (%)
phenolic
of 20 %;
compounds
HPCDAE (control) (at atmospheric
pressure and temperature of 80 ºC)
anthocyanins
phenolic
compounds
PLE (at 117 bar, temperature of 80 ºC,
anthocyanins
and flow rate of 1 mL of acidified
water/min).
phenolic
compounds
*n.d – not determinate
The increase in the extraction efficiency and in the mass transfer rates (M*),
which accelerates the extraction process of anthocyanins and other phenolic compounds, is
111
probably also associated to the solvent extraction viscosity modification, which can change
the solubilities of the desired compounds in the extraction solvent. The viscosity value in
this study varied from 0.35603 to 0.65424 cP (http://webbook.nist.gov/chemistry/fluid). In
agreement with previous studies (Ju & Howard, 2003; Cacace & Mazza, 2002) the decrease
of the viscosity coefficient increases the extraction efficiency and accelerates the extraction
process. HPCD Assisted-Extraction optimum conditions resulted in a water viscosity
coefficient (0.35741 cP) very low, which might produce an increase of the diffusion of the
analytes from the sample matrix into the extraction solvent. According to Yamaguchi &
Kimura (2000) a decrease of the viscosity coefficient significantly increases solutes
diffusivities.
As the extraction solvent of the proposed extraction method for anthocyanins
and other phenolic compounds synchronously is carbonated water with small quantities of
HCl, this emerging pressure extraction method may be considered a promising
environment-friendly technology for this class of bioactive compounds from other sources.
To our knowledge, there are rare studies about the use of HPCD system for bioactive
compounds extraction purpose.
6.4 Conclusions
Process variables had significant effect on the extraction of functional
components with extraction pressure, temperature and their interactions being significant
for the extraction of anthocyanins and extraction temperature being highly significant for
the extraction of phenolic compounds. The effects of extraction pressure, volume ratio of
solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) and the interaction between extraction
112
temperature and volume ratio of solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) were
also significant for the extraction of phenolic compounds.
The optimum HPCD Assisted-Extraction conditions for the extraction of the
bioactive compounds (anthocyanins and phenolic compounds synchronously) from
jabuticaba skins were determined by an optimization method that combined Response
Surface Methodology (RSM) with desirability function and the predicted values for the
recovery of these bioactive compounds were well consistent to the experimental ones. The
optimum conditions include 117 bar extraction pressure, 80 ºC extraction temperature and
20% volume ratio of solid–liquid mixture/pressurized CO2 ( R S−L / CO 2 (%) ) for the total
anthocyanin and phenolic compounds contents of 2.24 ± 0.31 mg cyanidin-3-glucoside/g
dry skins and 12.90 ± 1.46 mg gallic acid equivalents/g dry skins, respectively.
Compared to PLE and to control experiment (without pressurization) the
experimental results obtained using optimum HPCD Assisted-Extraction conditions was
much more effective and faster in extracting anthocyanins and phenolic compounds from
jabuticaba skins.
Extraction of anthocyanins and other phenolic compounds using carbonated
water by High Pressure CO2 with small amounts of HCl is probably related to: (1)
disruption of the vegetable matrix by rapid CO2 depressurization, (2) pH decrease that
occurs due to the generation of “in situ” carbonic acid and/or alkyl carbonic acid, when the
CO2 is solubilized in water, (3) reduction of the dielectric constant of the extraction solvent
into the range of intermediate-behavior solvents such as methanol or ethanol by modifying
pressure and temperature, and possibly with the formation of ions HCO3-3, and CO3-2 in
acidified water, (4) increase of anthocyanin and phenolic compounds solubilities, (5)
reduction of solvent viscosity, and (6) increase of diffusion rate of the analytes from the
vegetable matrix to the solvent.
113
Acknowledgements
The authors are grateful to CNPq for financial support. Diego T. Santos thanks
CNPq (141894/2009-1) for the doctorate fellowship.
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116
CAPÍTULO 7 - MICRONIZATION AND ENCAPSULATION OF FUNCTIONAL
PIGMENTS USING SUPERCRITICAL CARBON DIOXIDE
Diego T. Santos and M. Angela A. Meireles
Trabalho submetido ao periódico Journal of Food Process Engineering.
117
Key words
Particle Formation, Functional Pigments, Rapid Expansion of Supercritical Solution
(RESS), Sensitive Compounds, Supercritical Anti-Solvent (SAS)
Abstract
This research involves experimental studies of supercritical fluids-based micronization and
encapsulation processes exploiting both solvent and anti-solvent properties of supercritical
CO2 for diverse functional pigments, in order to asses the reliability of home-made
experimental apparatuses and to validate the experimental data obtained with these
equipments. Quercetin and β-carotene were used as model substances in the micronization
process via SAS (Supercritical Anti-solvent). Bixin-rich extract with PolyEthylene Glycol
(PEG) 10000 as encapsulant material were used in the encapsulation process via SAS,
while rutin and anthocyanin-rich extract with PEG 10000 and ethanol as co-solvent were
applied to the formation of polymeric microcapsules via RESS (Rapid Expansion of
Supercritical Solutions). The processing parameters and the levels used were based on
earlier results achieved from other research groups. Core material:encapsulant material
ratio, core material physical properties such as solubility in supercritical CO2 and in CO2 +
ethanol and viscosity were key parameters for these processes.
Practical Applications: The application of natural food colorants with relevant antioxidant
activities, such as carotenoids and flavonoids, as food additives in various food products is
seriously hampered by their fast degradation triggered by light, temperature, presence of
oxygen, insolubility in aqueous systems, low dispersibility, among others. Their particle
size reduction and/or their encapsulation have been successfully used to overcome all these
drawbacks. Supercritical fluids (SCFs) have become an attractive alternative due to be
considered environmentally friendly solvents. SCFs may be conveniently used for various
118
applications such as extraction, reactions, micronization, encapsulation, among others.
Studies showing the use of SCFs and/or the construction of apparatuses that use these green
solvents for solubility enhancement of functional pigments with poor water solubility
and/or avoiding degradation of these compounds are extremely important.
7.1 Introduction
Supercritical fluids (SCFs) have become an attractive alternative due to be
considered environmentally friendly solvents (Prado et al. 2010). The methods that use
supercritical fluids can be conveniently used for various applications such as extraction,
micronization, encapsulation, etc. (Santos and Meireles 2010).
In recent years, novel particle formation techniques utilizing supercritical fluids
have been developed in order to overcome some of the disadvantages of the conventional
techniques. Some of these drawbacks are: a) poor control of particle size and morphology;
b) degradation of thermo sensitive compounds; c) low encapsulation efficiency; d) low
precipitation yield. Additionally, the use of supercritical fluids as phase separating agents
has been intensively studied also to minimize the amount of potentially harmful residues in
the capsules (Santos and Meireles 2010).
According to Cocero and coworkers carbon dioxide (CO2) is the most commonly
solvent used for micronization and encapsulation purposes because the supercritical region
can be achieved at moderate pressures and temperatures (Tc = 304.2 K, Pc = 7.38 MPa);
therefore, working with supercritical CO2 it is possible to carry out processes at nearambient temperatures, avoiding the degradation of thermolabile substances (Cocero et al.
2009). The great interest in processes that use supercritical CO2 is the result of the easy
removal of it from the final product, non-toxicity and non-flammability (Pereira et al.
2007). For extraction purposes due to similar reasons supercritical CO2 also is the most
widely used solvent (Braga et al. 2007).
119
Currently there is a trend towards a healthier way of living, which includes a
growing awareness by consumers for what they eat and what benefits certain ingredients
have in maintaining good health. The application of pigments extracted from plants cells
with several biological activities as food additives in various formulations is seriously
hampered by their fast degradation triggered by light, temperature, presence of oxygen,
insolubility in aqueous systems, low dispersibility, among others (Özen et al. 2009; Mattea
et al. 2009). Their particle size reduction and/or their encapsulation in general is used to
overcome all these challenges (Suo et al. 2005; Mattea et al. 2009).
Several precipitation processes of micro to nano range of particles both in pure and
encapsulated forms using SCFs have been developed. These processes can be classified
according to the role of the supercritical fluid in the process: solvent [Rapid Expansion of
Supercritical Solutions (RESS)]; Supercritical Solvent Impregnation (SSI), solute [Particles
from Gas Saturated Solutions (PGSS)] or anti-solvent [Supercritical Anti-Solvent (SAS);
Supercritical Fluid Extraction of Emulsions (SFEE)] (Martín and Cocero 2008).
Recent developments in functional pigments particle formation and co-precipitation
with biodegradable polymers using supercritical carbon dioxide as solvent or anti-solvent
have been successfully employed. The reproducibility of these results using physically
different apparatuses and on both laboratory and semi-industrial scale is extremely
important. Thus, the present study involves experimental studies of SCF-based
micronization and encapsulation processes exploiting both solvent and anti-solvent
properties of supercritical CO2 for diverse functional pigments in order to asses the
reliability of home-made experimental apparatuses and to validate the experimental data
obtained with these equipments.
Based on the solubility of the compound in supercritical CO2, two different
approaches were applied. Quercetin, a plant pigment with antioxidant properties that
belongs to the genre known as flavonoids (Hertog et al. 1992); β-carotene, a type of
carotenoid used as natural colorants in food products that acts as precursor of Vitamin A
and as antioxidant (Priamo et al. 2010) were used as model substances in the micronization
process via SAS. Bixin-rich extract, an extract with high concentration of the thermolabile
120
carotenoid type bixin, widely used in the food, pharmacological and cosmetic industries
due to the intensity of its color (Silva et al. 2008) with a biopolymer as encapsulant
material were used in the encapsulation process via SAS. In the RESS process, rutin, a
yellow flavonoid pigment with strong antioxidant activity (Nishikawa et al. 2007) and
anthocyanin-rich extract, an extract with high content of a type of colorful flavonoids easily
susceptible to degradation that shows several health promoting benefits (Santos and
Meireles 2009), with a biopolymer and a co-solvent were applied to the formation of
polymeric microcapsules.
7.2 Materials and methods
7.2.1 Materials
Quercetin dihydrate with a minimum purity of 98 % was purchased from
Sigma–Aldrich (St. Louis, USA). β-carotene (minimum purity of 97 %) was purchased
from Fluka (Buchs, Switzerland).
Figures 7.2.1.1 and 7.2.1.2 show, respectively, the quercetin and β-carotene
crystals prior to subjecting them to the micronization process via SAS (using SC-CO2 as
anti-solvent). Both Scanning Eletronic Microscope (SEM) micrographs presented flake-like
crystals with mean width of 7.590 μm for quercetin and 3.288 μm for β-carotene.
Quercetin is soluble in ethyl acetate (EA); β-carotene is soluble in
dichloromethane (DCM); EA and DCM are both highly miscible in supercritical CO2
resulting in high volumetric expansion in SC-CO2. Therefore, EA and DCM (analytical
grade) purchased from Merck (Darmstadt, Germany) were used to prepare the solutions.
121
Figure 7.2.1.1 - SEM micrograph of unprocessed quercetin sample.
Figure 7.2.1.2 - SEM micrograph of unprocessed β-carotene sample.
122
Bixin-rich extract was obtained from annatto (Bixa orellana L.) seeds using
pure supercritical CO2 at 31 MPa and 333 K. Bixin-rich extract and PolyEthylene Glycol
(PEG) with a mean molecular weight of 10000 g/mol (melting point: 63–65 ºC) (Sigma–
Aldrich, Steinhein, Germany) are both soluble in DCM, hence this solvent was used to
prepare the solutions.
Quercetin and β-carotene micronization processes and encapsulation process of
bixin-rich extract in PEG were carried out employing carbon dioxide as anti-solvent due to
its very low solubility as a compressed fluid in the temperature and pressure ranges
investigated.
Rutin hydrate with a minimum purity of 95 % was purchased from Sigma–
Aldrich (St. Louis, USA). Anthocyanin-rich extract was obtained from jabuticaba
(Myrciaria cauliflora) skins using pressurized ethanol at 0.5 MPa and 353 K.
Figure 7.2.1.3 shows the rutin crystals prior to subjecting them to the
encapsulation process via RESS (using SC-CO2 as solvent), which are flake-like, bar-like
and agglomerated crystals with mean particle size of 9.254 μm.
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Figure 7.2.1.3 - SEM micrograph of unprocessed rutin sample.
Ethanol with analytical grade purchased from Ecibra (Santo Amaro, Brazil) was
used in the encapsulation process of rutin and anthocyanin-rich extract via RESS as cosolvent to enhance the solubility of the polymer in the supercritical CO2. Additionally,
ethanol was chosen also due its nonsolvent property to the produced polymeric
microcapsules, avoiding any agglomeration after encapsulation procedure. PEG with a
mean molecular weight of 10000 g/mol (melting point: 63–65 ◦C) (Sigma–Aldrich,
Steinhein, Germany) was used as encapsulant material.
Dry carbon dioxide, 99.9% purity (Gama Gases Especiais Ltda., Campinas,
Brazil) used in all experiments was supplied in the liquid phase.
For the construction of the standand curves of absorbance vs. concentration in
the solvent chloroform (Ecibra, Santo Amaro, Brazil), distilled water and hydrochloric
acid/potassium chloride buffer pH 1.0 were used for the determination of bixin-rich extract,
rutin and anthocyanin-rich encapsulation efficiencies, respectively.
7.2.2 Micronization process via SAS
The experiments for quercetin and β-carotene micronization via supercritical
fluid were performed in the home-made SAS equipment; a schematic diagram of which is
shown in Figure 7.2.2.1. Liquid CO2, anti-solvent, was fed from the cylinder through a
thermostatic bath (Marconi, MA-184, Piracicaba, Brazil) at -10 °C to ensure the
liquefaction of the fluid and to prevent cavitation, and then it was pumped by an air driven
liquid pump (Maximator Gmbh, PP 111, Zorge, Germany) to the high-pressure vessel
(volume of 500 mL; 6.8 cm internal diameter) via a nozzle. The nozzle consists in a 1/16 in.
tube (inner diameter (i.d.): 177.8 μm) for the solution, placed inside a 1/8 in. tube for the
CO2. Once the particle formation vessel reached steady state (temperature, pressure and
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CO2 flow rate), the solution was introduced into the vessel by a HPLC pump
(Thermoseparation Products, Model ConstaMetric 3200 P/F, Fremoni, USA) through the
coaxial annular passage of the atomizer. Pressure was controlled by a back pressure
regulator (Tesco, model nº 26-1761-24-161, ELK River, USA). The vessel temperature was
kept constant by a heating water bath (Marconi, MA 127BO, Piracicaba, Brazil). CO2 flow
rate was measured using a glass float rotameter (0.15-2.2 kg/h of CO2 at 1.013bar/20 ºC,
ABB, 16/286A/2, Warminster, USA) coupled to a flow totalizer (LAO, model G0,6,
Osasco, Brazil). When the desired amount of solution (quercetin in ethyl acetate or βcarotene in dichloromethane) had been injected, which enabled the collection of sufficient
amount of precipitated powder for analysis, the HPLC pump was stopped and only pure
CO2 was fed. The flow of CO2 was maintained during 20 minutes for the complete removal
of solvent from the precipitator, which was proved necessary by the preliminary
experimental results. Quercetin and β-carotene precipitates were trapped by a paper filter
fixed at the bottom of the vessel, while the fluid mixture (SC-CO2 plus solvent) exited the
vessel and flowed to a second vessel (100 mL glass flask) connected after the micrometric
valve used to collect the particles carried with the effluent solution that leaves the
precipitation vessel. At the end, the precipitation vessel was slowly depressurized to
atmospheric pressure and particles were collected and stored in the dark in a domestic
freezer (-10 ºC) (Double Action, Metalfrio, São Paulo, Brazil) until subsequent analysis and
characterization. A heating system mantained at 120 ºC was used to heat the micrometric
valve to avoid Joule-Thomson freezing effect that can lead to clogging of the throttling
device during particle formation procedure.
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4
3
4
2
3
8
9
7
3
17
5
13
4
4
3
6
16
1
13
10
4
14
Solution
CO2
11
CO2
15
12
Figure 7.2.2.1 - Schematic diagram of the SAS apparatus. 1 CO2 cylinder; 2 CO2 filter; 3
manometers; 4 valves; 5 thermostatic bath; 6 CO2 pump; 7 back pressure regulator; 8
solution reservoir; 9 solution pump; 10 thermocouple; 11 precipitation vessel; 12 heating
bath; 13 temperature controllers; 14 micrometric valve with a heating system; 15 glass
flask; 16 glass float rotameter; 17 flow totalizer.
For quecetin micronization the flow rate of CO2 and quercetin solution (1.4
mg.mL-1) were 0.6 kg.h-1 and 0.2 mL.min-1, respectively, and the precipitation pressure and
temperature were 10 MPa 313.15 K, respectively. Conventional micronization of quercetin
was also carried out, which was used as a reference process to compare with the
supercritical fluid technique. Ten milliliters of quercetin in ethyl acetate solution (1.4
mg.mL-1) was prepared and then the solvent was evaporated using a rotary evaporator
(Laborota, model 4001, Vertrieb, Germany), with vacuum control (Heidolph Instruments
126
Gmbh, Vertrieb, Germany) and thermostatic bath at 313.15 K. The quercetin precipitates
were collected from the precipitation vessel (50 mL glass flask) and stored in the dark in a
domestic freezer (-10 ºC) (Double Action, Metalfrio, São Paulo, Brazil) until subsequent
analysis and characterization.
For β-carotene micronization the flow rate of CO2 and β-carotene solution (8
mg.mL-1) were 1.5 kg.h-1 and 1 mL.min-1, respectively, and the precipitation pressure and
temperature were 0.8 MPa 313.15 K, respectively.
The experimental conditions and procedures applied is this study were
employed in order to compare the results obtained in this work with the results obtained by
other research groups (Can et al. 2009; Franceschi et al. 2009) at the same operational
conditions using similar apparatuses. All experiments were done in duplicate.
7.2.3 Encapsulation process via SAS
The experiments for bixin-rich extract encapsulation in PEG via supercritical
fluid were performed in the SAS equipment described before. The encapsulation procedure
was very similar to the micronization procedure described differing that an encapsulant
material was added to the solution containing the core material dissolved in a solvent. PEG
in this case was the encapsulant material, Bixin-rich extract the core material and
dichloromethane the solvent. Dichloromethane was the selected solvent because it is a good
solvent for bixin-rich extract and PEG.
The solution flow rate, precipitation pressure and temperature were fixed at 1
mL.min-1, 10 MPa and 313.15 K, respectively. The CO2 flow rate and mass ratio between
127
bixin-rich extract and PEG investigated were 0.6 and 1.5 kg.h-1; 1:2 and 1:10, respectively.
All experiments were done in duplicate.
7.2.4 Encapsulation process via RESS
A schematic diagram of the home-made RESS equipment is given in Figure
7.2.4.1. The RESS equipment consists of a CO2 pump (Maximator Gmbh, PP 111, Zorge,
Germany), a stainless steel pre-expansion vessel (6.57 mL, Thar Designs, Pittsburg, USA)
containing syntherized metal filters at the inlet and outlet, a spray nozzle and an expansion
vessel. A certain amount of core material (rutin or anthocyanin-rich extract), encapsulant
material (PEG) and co-solvent (ethanol) was first loaded into the pre-expansion vessel.
After being carefully sealed, liquid carbon dioxide cooled to around -10 °C by a
thermostatic bath (Marconi, MA-184, Piracicaba, Brazil) was delivered into the preexpansion vessel. Simultaneously, with the addition and pressurization of CO2 the preexpansion vessel was heated by an electric jacket until the desired supercritical conditions
were achieved. Pressure was controlled by a back pressure regulator (Tesco, model nº 261761-24-161, ELK River, USA). After achieving equilibrium, the mixture was kept for
about 30 min to ensure that the core and coating materials were dissolved in the
supercritical CO2. The prepared supercritical solution (a solution or suspension of core
material in a CO2-containing co-solvent and the dissolved polymer) was then sprayed
through a stainless steel capillary nozzle for a short time (< 4 s) to atmospheric pressure by
the opening of the valve placed before the nozzle. The nozzle consists in a 1/16 in. tube
(inner diameter (i.d.): 0.51 mm). The nozzle was maintained at 333 K with an electric
heater to allow its rapid expansion avoiding Joule-Thomson freezing effect that can lead to
128
clogging of the throttling device during the rapid expansion. The distance from the orifice
exit to the expansion target (the internal wall of the expansion vessel) was approximately 3
cm.
3
4
2
3
7
3
4
5
4
6
4
10
8
1
4
9
11
12
Figure 7.2.4.1 - Schematic diagram of the RESS apparatus. 1 CO2 cylinder; 2 CO2 filter; 3
manometers; 4 valves; 5 thermostatic bath; 6 CO2 pump; 7 back pressure regulator; 8 preexpansion vessel; 9 micrometric valve with a heating system; 10 temperature controller; 11
nozzle; 12 expansion vessel.
The residual solvent (ethanol) in the particles was measured by the recording of
the weight loss after the heating of a known amount of sample at 363.15 K for 24 hours.
Based on the results of previous works (Matsuyama et al. 2003; Kongsombut et
al. 2009) using similar apparatuses, core material and encapsulant material the
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experimental conditions were selected. For both encapsulation processes the same
operational conditions were employed: pre-expansion pressure and temperature of 20 MPa
and 313.15 K, ethanol and PEG concentration of 27.1 % (w/w) and 8.1 % (w/w),
respectively. Two mass ratios between core material and PEG of 1:2 and 1:10 were used for
rutin encapsulation process. The amount of CO2 required for filling the RESS system is
8.43 g. All experiments were done in duplicate.
7.2.5 Characterization and Analysis
7.2.5.1 Particle characterization
Micronized and encapsulated particles were analyzed by Scanning Electron
Microscopy (SEM) (LEO 440i, Leica, Cambridge, USA) after coated with a thin gold film
with the aid of a sputter coater (Polaron, SC 7620, Ringmer, England) to determine particle
morphology, particle size and particle size distribution.
The particle size distribution was obtained according to the following
procedures. Initially, with the aid of a software (Micro Image Analysis Software, version
2.2, Eletronic Eyepiece, Zhejiang, China), 100 randomly selected, well-separated particles
from the SEM images were measured in zoom-in mode, in which individual particles can
be recognized clearly. Secondly, the particle size was calculated based on the ratio of their
width or length to the SEM magnification scale. And finally, a histogram for particle size
distribution was drawn, the mean particle size, PS (μm), and the standard deviation, SD
(μm), in normal distribution mode N (PS, SD) were estimated. Other two ways to
characterize the size distribution of the produced particles were also used alternatively: (1)
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by plotting the particle size of the particles as cumulative distribution; (2) by calculating the
variation coefficient (VC) by the Equation 1:
VC (%) =
s tan dard deviation ( SD)
X 100
mean particle size ( PS )
(1)
The color of PEG is obviously different from that of core materials, so the
characteristics of dispersion of the core materials (bixin-rich extract, rutin and anthocyaninrich extract) in the polymeric encapsulant material was analyzed by color difference
between them using optical micrographs (Nikon, mod Eclipse E200, Tokyo, Japan).
7.2.5.1.1 Determination of Precipitation Yield - PY (%)
The precipitation yield – PY (%) was evaluated considering the amount of
micronized powder collected in the precipitation vessel. The percentage of precipitation
yield was calculated by the ratio between the mass of quercetin and β-carotene collected in
the precipitation vessel after each assay and the mass of them present in the organic
solution added to the precipitation vessel at each experiment.
7.2.5.1.2 Determination of Encapsulation Efficiency (EE (%))
The encapsulation efficiency was verified by a UV–Vis spectrophotometer
(Hitachi, model U-3010, Tokyo, Japan). First, a sample of core material encapsulated in
PEG was weighed. It was assumed that the ratio between core material and encapsulant
material remained constant after the precipitation. Afterwards, the sample was dissolved in
a suitable solvent for both substances (core material and PEG) and the absorbance was
measured at maximum absorbance wavelengths. For bixin-rich extract, rutin and
anthocyanin-rich extract different solvents were used: dichloromethane, distilled water and
hydrochloric acid/potassium chloride buffer pH 1.0, and at different wavelengths 438, 352
and 512 nm, respectively. Comparing the results with a standard curve of absorbance vs.
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concentration of free core material in the solvent, the amount of bixin-rich extract, rutin and
anthocyanin-rich extract encapsulated were evaluated by a straight forward calculation of
the encapsulation efficiency (EE (%)) by the Equation 2:
EE (%) =
mass of core material encapsulated
X 100
theoretical mass of core material encapsulated
(2)
7.3 Results and discussion
7.3.1 Micronization process via SAS
7.3.1.1 Quercetin micronization
In Figure 7.3.1.1.1, the SEM micrograph shows the change in morphology and
size of quercetin particles obtained using our home-made SAS equipment. The precipitates
obtained under the experimental condition employed were needle-like particles with mean
length of 1.872 μm. Comparing the sizes of micronized particles by SAS with that of
unprocessed quercetin crystals (Figure 7.2.1.1), it can be concluded that the SAS equipment
used in this study could be used to make micronized quercetin particles. In this study, the
particle size from micronized quercetin by SAS was 4.1 times smaller than unprocessed
quercetin.
Quercetin was also micronized by conventional solvent evaporation method to
obtain the reference materials for comparison. SEM micrograph of the micronized
quercetin is shown in Figure 7.3.1.1.2. It was not observed a change in particle
morphology. The morphology of flake-like of the quercetin crystals remained after
conventional micronization process. On the other hand, the particle size from micronized
quercetin by this technique was only 1.8 times smaller than unprocessed quercetin.
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Figure 7.3.1.1.1 - SEM micrograph of quercetin micronized particles obtained by SAS
process.
Figure 7.3.1.1.2 - SEM micrograph of quercetin micronized particles obtained by
conventional process.
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Similar results were observed by Can et al. (2009) using a similar SAS
apparatus for quercetin micronization. A significant reduction of the quercetin particles (3.2
times) was also observed under the same experimental condition employed in this study,
though no change in morphology occurred to quercetin particles that remained at their
initial form. The slightly difference (21.1 %) between the size reduction results can be
associated mainly to two factors: (1) the inner diameter of the coaxial annular passage of
the nozzle; (2) the size and geometry characteristics of the precipitation vessel. Since no
information about these factors is provided by the authors, no conclusions can be inferred.
Moreover, the same study also shows that quercetin micronized particles made by SAS
process is much smaller than those obtained by conventional solvent evaporation method. It
was suggested by the authors that the reason may be that at high velocity of supercritical
CO2 the solution is broken into very small droplets in the SAS process, resulting in the
formation of fine particles.
Regarding the size distribution of the micronized particles, Figure 7.3.1.1.3
shows in terms of cumulative distribution the size distribution of the quercetin particles
micronized by SAS and by conventional technique. The technique employed had no
significant effect on the particle size distribution of the micronized precipitates. On the
other hand, the technique employed has influenced the precipitation yield - PY (%).
Different PYs, depending on the experimental method were obtained. A higher PY (99.5
%) was obtained using conventional micronization process than using SAS process (81.9
%).
Franceschi et al. (2009) found comparable precipitation yields (in the range of
71-94 %) depending on the process parameter values for β-carotene micronization via SAS.
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Nevertheless, other previous studies have obtained higher PYs for diverse particles. Miguel
et al. (2008) for lutein micronization from also ethyl acetate solutions via SAS have
obtained precipitation yields above 95 %. The amount of lutein precipitated and the
precipitation yield were calculated by Miguel and coauthors as the difference between the
amount of lutein in the feed, and the amount of lutein collected in the liquid effluent, while
in our study and in the study of Franceschi and coauthors a direct calculation with the
weight of the product in the precipitation vessel was done. As the last procedure seems to
be greatly influenced by the collection efficiency of the particles, probably the lower value
Cumulative distribution (%)
of PY observed in this study is mainly related to the procedure employed.
100
90
80
70
60
50
40
30
20
10
0
0,1
1
10
100
1000
Particle size (μm)
Micronized particles by SAS
Micronized particles by conventional process
Figure 7.3.1.1.3 - Size distribution of quercetin particles obtained by SAS and conventional
processes.
7.3.1.2 β-carotene micronization
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The morphology of β-carotene after SAS process as occurred with quercetin
also changed. The unprocessed β-carotene presented a flake-like form (Figure 7.2.1.2),
while the precipitated β-carotene a leaf-like form (Figure 7.3.1.2.1). Moreover, a change in
the size of particles obtained by SAS process was also observed. An increase in the particle
size was observed: starting from unprocessed β-carotene with mean particle size of 3.288
μm, the mean particle size of precipitated β-carotene was 16.090 μm.
Figure 7.3.1.2.1 - SEM micrograph of β-carotene micronized particles obtained by SAS
process.
Although, the objective of reducing β-carotene particles was not achieved, the
results obtained in this work seem to be consistent with current scientific literature.
Franceschi et al. (2008) noted that most of the experimental runs of their work produced
larger particles, in a wide size range dependent on the process conditions used, compared to
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the unprocessed β-carotene particles. Using a different precipitation vessel in another study
for micronization of β-carotene from dichloromethane solutions as well, Franceschi and
coauthors observed a reduction in the mean particle size at the same operational conditions
(Franceschi et al. 2009). It was suggested that the verified behavior might be explained by
the enlargement of the precipitation vessel. In the first work, the inner diameter and height
of the precipitation vessel were smaller than of the one used in the second work (resulting
in a much smaller volume), thus the larger precipitation vessel possibly allowed an efficient
removal of the solvent before particle deposition in the bottom or walls of the vessel,
producing smaller particles. Since the precipitation vessel used in this study is slightly
smaller than of the used by Francheschi and coauthors in the second work (16.7 %), it
might be affected the recrystalization of β-carotene via SAS process.
Concerning the inner diameter of the coaxial annular passage, a size decrease
leads to a particle size decrease. According to Boutin et al. (2009) a smaller inner diameter
favors the organic solution dispersion (particularly by increasing the injection speed for the
same flow rate) and therefore the formation of smaller particles. In this way, the large
difference between the precipitated β-carotene size obtained in our study (16.090 μm) and
the obtained by Francheschi and coauthors (3.2 μm) using the same processing conditions
might be highly linked to the much lower inner diameter of the coaxial annular passage
used in by Francheschi et al. (2009) (100 μm).
From these research results it is suggested that the combination of the use of a
smaller precipitation vessel (500 mL) and a larger inner diameter of the coaxial annular
passage (177.8 μm) produced larger β-carotene particle size.
137
Comparing the morphology of β-carotene precipitated particles obtained in this
work with that obtained by Francheschi et al. (2009) at the same operational conditions, it
was observed a strong similarity. Both of them presented leaf-like forms. It was
demonstrated that the particle morphology is extremely dependent on the operational
process conditions employed, being the justification for obtaining leaf-like β-carotene form
by SAS process is the high anti-solvent flow rate used. The use of high CO2 flow rate
causes a pronounced turbulence inside the precipitation vessel leading to an increase in the
kinetic energy of the atomizing CO2. Thus, the mass transfer rates between CO2 and the
organic solvent are increased; with CO2 diffusing more rapidly into the droplet and the
solvent evaporating from droplets instantaneously causing accelerated supersaturation and
nucleation (Francheschi et al. 2008; 2009), which might affect the recrystallization process
generating a modification of the particle morphology.
Likewise, our results on β-carotene precipitation yield (87.7 %) are in
agreement with those of Francheschi et al. (2009) (87.0 %).
7.3.2 Encapsulation process via SAS
7.3.2.1 Bixin-rich extract encapsulation
Two different CO2 flow rates were investigated. The higher CO2 flow rate
employed resulted in an excessive loss of particles. It was visually observed in the
downstream connections of the precipitation vessel and in the 100 mL glass flask the
presence of bixin-rich extract. Since the bixin-rich extract was produced using supercritical
CO2 at 31 MPa and 333 K, during SAS process part of the extract, which is soluble in CO2
at 10 MPa and 313 K, was carried with the effluent solution (CO2 plus dichloromethane).
138
Using the lower CO2 flow rate the loss of bixin-rich extract encapsulated was reduced, thus
the lower CO2 flow rate (0.6 kg.h-1) would be preferred for further experiments.
The effect of mass ratio between bixin-rich extract and PEG on the
encapsulation process was also evaluated. According to our findings and corroborating the
literature data, a decrease in the mass ratio between core material and encapsulant material
led to the production of less agglomerated particles (Martín et al. 2007). Then, the smaller
(1:10) mass ratio between bixin-rich extract and PEG was chosen. For the mass ratio of 1:2
probably the amount of polymer was not sufficient to effectively encapsulate the amount of
bixin-rich extract.
Microparticles of polymer PEG with other carotenoids loaded within have been
successfully produced using supercritical CO2 as anti-solvent, such as β-carotene (Martín et
al. 2007; He et al. 2007; Mattea et al. 2008) and lutein (Martín et al. 2007).
In agreement with those studies, bixin-rich extract was loaded into the PEG
matrix with success. Unprocessed PEG powders, demonstrated in Figure 7.3.2.1.1-1, are
gray and transparent slice-like particles with sizes about 2–5 mm. With color difference
between bixin-rich extract (orange) and PEG there is a clear evidence of the presence of
well dispersed bixin-rich extract inside the polymer matrix obtained by SAS process (CO2
flow rate of 0.6 kg.h-1; mass ratio between core material and PEG of 1:10) (Figure
7.3.2.1.1-2).
The SEM micrograph (Figure 7.3.2.1.2-1) shows the morphology of the
microparticles of bixin-rich extract encapsulated in PEG. Diverse morphologies were
observed, such as flake-like, bar-like, etc. He et al. (2006) also produced β-carotene/PEG
particles using similar SAS equipment, and verified a variety of morphologies.
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Figure 7.3.2.1.1 - Optical micrographs of: 1) PEG sample; 2) bixin-rich extract
encapsulated in PEG by SAS process (CO2 flow rate of 0.6 kg.h-1; mass ratio between core
material and PEG of 1:10); 3) rutin encapsulated in PEG by RESS process (mass ratio
between core material and PEG of 1:10); 4) anthocyanin-rich extract encapsulated in PEG
by RESS process (mass ratio between core material and PEG of 1:10).
Bixin-rich extract/PEG particle size of 33.02 ± 2.17 µm was produced in this
study. Mattea et al. (2008) employing similar SAS processing conditions produced βcarotene/PEG particle sizes in the range of 15-200 μm, corroborating our results.
140
Figure 7.3.2.1.2 - SEM micrographs of: 1) bixin-rich extract encapsulated in PEG by SAS
process (CO2 flow rate of 0.6 kg.h-1; mass ratio between core material and PEG of 1:10); 2)
rutin encapsulated in PEG by RESS process (mass ratio between core material and PEG of
1:2); 3) rutin encapsulated in PEG by RESS process (mass ratio between core material and
PEG of 1:10); 4) anthocyanin-rich extract encapsulated in PEG by RESS process (mass
ratio between core material and PEG of 1:10).
The encapsulation efficiency (EE (%)) for bixin-rich extract in PEG under the
SAS condition investigated in this work was determined and compared to literature data.
An EE value of 62.23 ± 4.32 % was obtained. Encapsulation efficiencies ranging around 5141
80 % have been reported for other functional pigments, depending on SAS processing
conditions, such as mass ratio between core material and encapsulant material, precipitation
pressure and temperature, etc. (He et al. 2006; Francheschi et al. 2009). Furthermore, the
procedure for the determination of the EE may influence its value. The procedure that takes
into account the minor amount of core material adsorbed onto the polymer surface results
obviously in a EE slightly higher than the EE determinated using a procedure that does not
do this consideration (Priamo et al. 2010). Due to the simplicity of the last procedure we
decided to use it in this work.
7.3.3 Encapsulation process via RESS
7.3.3.1 Rutin encapsulation
The Rutin/PEG particles (Figures 7.3.2.1.2-2; 7.3.2.1.2-3) had an apparently
amorphous morphology, very different from the crystalline morphology of rutin crystals
prior to subjecting them to the encapsulation process (Figure 7.2.1.3), indicating that rutin
particles had been successfully encapsulated in PEG matrix using our home-made RESS
equipment using ethanol as co-solvent.
The effect of mass ratio between rutin and PEG on the encapsulation efficiency
(EE (%)) and rutin/PEG particle size was investigated. For the mass ratio between rutin and
PEG of 1:2 the encapsulation efficiency was 44.18 ± 0.59, while for the mass ratio of 1:10
the EE was 21.19 ± 2.67. A decrease in the concentration of core material resulted in a
decrease in the encapsulation efficiency and in the particle size (Figures 7.3.2.1.2-2;
7.3.2.1.2-3), keeping fixed PEG content, as expected. Whereas
Matsuyama et al. (2003)
using similar RESS apparatus, core material and experimental conditions have not
142
estimated the EE values for their microcapsules it was suggested that most of the feed
flavone particles were well coated with PEG being mainly located in the center of the
polymeric microcapsules. Figure 7.3.2.1.1-3 shows that the rutin particles (mass ratio of
1:10) were well distributed in the microcapsules, corroborating the findings of
Matsuyama’s research group.
According to Matsuyama et al. (2003) the key of the encapsulation via RESS
using co-solvents is the ability to control the thickness and particle size distribution of the
microcapsules with the feed concentration of the polymer. Since the mean particle size of
the rutin/PEG obtained in this study was 42.944 µm (mass ratio of 1:10), while of
unprocessed rutin particles was 9.254 µm, it can be suggested that, or there is a large
thickness of PEG on each rutin particle, or each rutin/PEG particle is an agglomerate of
rutin and PEG particles.
Based on the Kongsombut et al. (2009) theory is suggested that probably the
flow turbulence and friction loss generated during the rapid expansion of supercritical
solution process would not be sufficient to disintegrate all of agglomerates into totally
separated particles. In contrast, after expansion as expected the produced rutin/PEG
particles did not tend to agglomerate because the residual co-solvent (ethanol) acts as a
good nonsolvent for the polymer. The amount of residual ethanol in the microcapsules was
0.40 % (w/w). Matsuyama et al. (2003) obtained an amount of residual ethanol in their
microcapsules less than 1 % (w/w) for different polymers.
Regarding the size distribution of the rutin/PEG particles, due to the
experimental discrepancies observed for particle size and particle distribution, it was
decided to characterize the size distribution by the variation coefficient (VC). Several
143
research groups have employing the same criterion to determine the dispersion of particle
size in the precipitation using supercritical CO2 (Tenório et al. 2007; Franceschi et al. 2008;
2009; Priamo et al. 2010). For the mass ratio between rutin and PEG of 1:2 the variation
coefficient (VC) was 10.93 %, while for the mass ratio of 1:10 the VC was 22.85 %.
Depending on the application of the produced particles it will be required a process that can
produce microparticles with a desired mean particle size with an acceptable particle size
dispersion and with a significant core material encapsulated (high encapsulation
efficiency), hence it is extremely important the determination of all this response variables
for optimization studies.
7.3.3.2 Anthocyanin-rich extract encapsulation
The anthocyanin-rich extract/PEG particles (Figure 7.3.2.1.2-4) had also an
apparently amorphous morphology, indicating that anthocyanin-rich extract had been
successfully encapsulated in PEG matrix via RESS using ethanol as co-solvent.
With color difference between anthocyanin-rich extract (purple) and PEG (gray
and transparent) it was confirmed that anthocyanin-rich extract is located inside the
polymer matrix obtained by RESS process (mass ratio between core material and PEG of
1:10) (Figure 7.3.2.1.1-4), but its dispersion into the PEG matrix was not good using the
operational conditions used in this study.
Vatai et al. (2009) and Seabra et al. (2010) have demonstrated that anthocyanin
pigments have high solubility in supercritical CO2-ethanol. Given that anthocyanins
possibly have higher solubility in the mixture supercritical CO2 + ethanol than rutin due to
the attached sugar moiety of the anthocyanin molecule (Santos et al. 2010), when rutin was
employed as core material a process more similar to a rapid expansion of a suspension of
144
supercritical CO2-insoluble particles in the supercritical CO2 solution was carried out.
According to Tsutsumi et al. (2003) in this process the core material is not dissolved but
suspended in a single homogeneous mixture of CO2, co-solvent, and the dissolved
encapsulant material, leading to a homogeneous deposition of the encapsulant material on
the surface of the suspended particles, thereby generating a polymer encapsulating layer on
the particle surfaces.
Kongsombut et al. (2009) using SiO2 particles concluded that the dispersion
and segregation of SiO2 powder contributed to the low agglomeration tendency of the
encapsulated particles. Since that anthocyanin-rich extract is a thick and viscous extract and
not a powder as rutin and SiO2, it would be expected that anthocyanin-rich extract/PEG
particles would have a higher agglomeration tendency than rutin/PEG particles. In
agreement with this explanation Figure 7.3.2.1.2-4 shows that anthocyanin-rich
extract/PEG particles in fact have higher agglomeration tendency than rutin/PEG particles
(Figures 7.3.2.1.2-2; 7.3.2.1.2-3).
On the contrary, the encapsulation efficiency of anthocyanin-rich extract in
PEG matrix (49.42 ± 2.32 %) was higher than that using rutin as core material, possibly due
to its sticky characteristic. On the other hand, the amount of the residual ethanol in these
microcapsules and the variation coefficient were comparable to rutin microcapsules 0.52 %
(w/w) and 11.989 %, respectively.
7.4 Conclusions
This work investigated the micronization and encapsulation of diverse
functional pigments using supercritical carbon dioxide as solvent or anti-solvent.
145
Experimental results on functional pigments micronization and encapsulation already
studied using similar apparatuses were provided demonstrating the reliability of our homemade systems.
SAS process successfully reduced the particle size of quercetin by 4.1 times,
while conventional micronization process only 1.8 times. Although the objective of
reducing β-carotene particles was not achieved the results obtained in this work seem to be
consistent with current scientific literature, validating our SAS equipment. Furthermore, it
was demonstrated that SAS process can be successfully utilized to co-precipitate
microparticles of PEG loaded with bixin-rich extract. The results allow to identify CO2
flow rate, core material:encapsulant material ratio and core material solubility in
supercritical CO2 as the key parameters of the co-precipitation. Further experiments will be
carried out in order to optimize the processing parameters.
RESS process using ethanol as co-solvent was effectively employed to
encapsulate rutin and anthocyanin-rich extract in PEG matrix. The data obtained in this
study are in good agreement with the previous values reported by several authors using
similar operational conditions and equipments. Core material:encapsulant material ratio and
core material physical properties such as solubility in supercritical CO2 + ethanol and
viscosity were key parameters for this process. In order to better understand this
encapsulation technique further experiments will be done.
This work is part of a broader project aiming at developing alternative
sustainable technologies towards particle formation of sensitive bioactive compounds from
vegetable sources, hence the results obtained here may be relevant as a support for the
146
successful design of particles with the desired size, form and morphology, which can be
applied mainly in food products.
Acknowledgements
The authors are grateful to CNPq for financial support. Diego T. Santos thanks
CNPq (141894/2009-1) for the doctorate fellowship.
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151
152
CAPÍTULO 8 - STABILIZATION OF ANTHOCYANIN EXTRACT FROM
JABUTICABA SKINS BY ENCAPSULATION USING SUPERCRITICAL CO2 AS
SOLVENT
Diego T. Santos, Juliana Q. Albarelli, Marisa M. Beppu and M. Angela A. Meireles
Trabalho submetido ao periódico Industrial & Engineering Chemistry Research.
153
Key words
Anthocyanins, Encapsulation, Rapid Expansion of Supercritical Solution (RESS), Caalginate Beads
Abstract
Supercritical fluids (SCFs) have become an attractive alternative because they are
considered environmentally friendly solvents. In the recent years, novel particle formation
techniques utilizing supercritical fluids have been developed in order to overcome some of
the disadvantages of the conventional techniques. In this context, the present study
evaluates the stabilization of anthocyanin extract obtained from jabuticaba (Myrciaria
cauliflora) skins in Polyethileneglicol using supercritical CO2 as solvent and ethanol as cosolvent. For comparison, a conventional method called ionic gelification was employed to
produce encapsulated particles by entrapment in Ca-alginate beads.
8.1 Introduction
Anthocyanins are one of the most important groups of water-soluble and
vacuolar pigments in nature. They belong to the flavonoid group responsible for a wide
variety of colors of fruits, flowers and leaves ranging from salmon pink through red and
violet to dark blue1. This pigment has attracted great interest due to the wide range of
biological
activities
including
antioxidant2,
anti-inflammatory3,4,
anticancer5,
antimutagenic6 and chemopreventive activities7.
However, the introduction of anthocyanins into food and/or medical field has
proved to be a major technological challenge since these compounds have low stability to
the environmental conditions during processing, storage and also consumption. They are
susceptible to degradation through factors such as light, pH, temperature, presence of
154
sulfite, ascorbic acid, enzymes, among others8. One possible way to effectively protect
these compounds, from product processing to consumption, could be the use of
encapsulation techniques.
Encapsulation facilitates light- and heat-labile molecules to maintain their
stability and improve their shelf lives and effects. It is a rapidly expanding technology,
highly specific, with affordable costs9,10. Among diverse encapsulation techniques
available, only few were evaluated for anthocyanin encapsulation as spray drying using
different coating materials11, gelation using polymers as sodium alginate, pectin, curdlan12
and glucan13 and liophilization14.
Supercritical fluids (SCFs) have become an attractive alternative because they
are considered environmentally friendly solvents. In the recent years, novel particle
formation techniques utilizing supercritical fluids have been developed in order to
overcome some of the disadvantages of the conventional techniques. Some of these
drawbacks are: a) poor control of particle size and morphology; b) degradation and lost of
biological activity of thermo sensitive compounds; c) low encapsulation efficiency; d) low
precipitation yield. Additionally, the use of supercritical fluids as phase separating agents
has been intensively studied also to minimize the amount of potentially harmful residues in
the capsules10, 15.
In this context, the present study evaluates the encapsulation of anthocyanins
extracted from jabuticaba (Myrciaria cauliflora) skins, a Brazilian potential source of
anthocyanins1, using supercritical CO2 as solvent, ethanol as co-solvent and
Polyethileneglicol as encapsulant material in order to increase and retain this pigment
stability, protecting these compounds from environmental conditions and therefore
promoting a larger utilization of these natural compounds in industry. For anthocyanin
extract stabilization comparison, a conventional method called ionic gelification was
employed to produce encapsulated particles by entrapment in Ca-alginate beads.
8.2 Materials and methods
155
8.2.1 Plant Material
Jabuticaba fruits (Myrciaria cauliflora) harvested from a plantation in the State
of São Paulo, Brazil, were acquired from a fruit and vegetable market centre (CEASACampinas, Brazil). Immediately after acquiring, the fruits were stored in the dark in a
domestic freezer (-10 ºC) (Double Action, Metalfrio, São Paulo, Brazil) until sample
preparation. Before extraction, the fruits were manually peeled.
8.2.2 Anthocyanin Extraction
For the anthocyanin extraction Pressurized Liquid Extraction process was used.
Based on our previous experiments extraction pressure, temperature, static extraction time
and ethanol flow rate were set at 50 bar, 80 ºC, 9 min and 1.67 mL/min (during 12 min).
After PLE, anthocyanin extracts were rapidly cooled to 5 °C in ice water to prevent
anthocyanin degradation. Subsequently, the extraction cell was exaustively purged with a
flow rate of 0.71 kg/h of carbon dioxide 99.9% (Gama Gases Especiais Ltda., Campinas,
Brazil) during 8-9 min to ensure that no residual anthocyanin extract solution would be into
the extraction cell. At the end, ethanol from the extract solution was evaporated using a
rotary evaporator (Laborota, model 4001, Vertrieb, Germany), with vacuum control
(Heidolph Instruments Gmbh, Vertrieb, Germany) and thermostatic bath at 40 °C. All the
extracts were stored (- 10 ºC) in the dark until being used in the encapsulation processes.
8.2.3 Encapsulation of Anthocyanin Extract
8.2.3.1 By Rapid Expansion of Supercritical Solution (RESS) process
A schematic diagram of the home-made RESS equipment is given in Figure
8.8.2.3.1.1. The RESS equipment consists of a CO2 pump (Maximator Gmbh, PP 111,
156
Zorge, Germany), a stainless steel pre-expansion vessel (6.57 mL, Thar Designs, Pittsburg,
USA) containing syntherized metal filters at the inlet and outlet, a spray nozzle and an
expansion vessel. A certain amount of anthocyanin extract, encapsulant material
(Polyethylene glycol) and co-solvent (ethanol) were first loaded into the pre-expansion
vessel. Polyethyleneglycol (PEG) with a mean molecular weight of 10000 g/mol (melting
point: 63–65 ◦C) (Sigma–Aldrich, Steinhein, Germany) was used as encapsulant material.
Ethanol was used in the encapsulation process as co-solvent to enhance the solubility of the
polymer in the supercritical CO2. Additionally, ethanol was chosen due its nonsolvent
property to the produced polymeric microcapsules, avoiding any agglomeration after
encapsulation procedure. After being carefully sealed, liquid carbon dioxide (99.9% purity,
Gama Gases Especiais Ltda., Campinas, Brazil) cooled to around -10 °C by a thermostatic
bath (Marconi, MA-184, Piracicaba, Brazil) was delivered into the pre-expansion vessel.
Simultaneously, with the addition and pressurization of CO2 the pre-expansion vessel was
heated by an electric jacket until the desired supercritical conditions were achieved.
Pressure was controlled by a back pressure regulator (Tesco, model nº 26-1761-24-161,
ELK River, USA). After achieving equilibrium, the mixture was kept under established
condition for about 30 min to ensure that the core and coating materials were dissolved in
the supercritical CO2. The prepared supercritical solution (a solution or suspension of core
material in a CO2-containing co-solvent and the dissolved polymer) was then sprayed
through a stainless steel capillary nozzle for a short time (< 4 s) to atmospheric pressure by
the opening of the valve placed before the nozzle. The nozzle consists in a 1/16 in. tube
(inner diameter (i.d.): 0.51 mm). The nozzle was maintained at 333 K with an electric
heater to allow its rapid expansion avoiding Joule-Thomson freezing effect that can lead to
157
clogging of the throttling device during the rapid expansion. The distance from the orifice
exit to the expansion target (the internal wall of the expansion vessel) was approximately 3
cm.
3
4
2
3
7
3
4
5
4
6
1
4
10
8
4
9
11
12
Figure 8.8.2.3.1.1 - Schematic diagram of the RESS apparatus. 1 CO2 cylinder; 2 CO2
filter; 3 manometers; 4 valves; 5 thermostatic bath; 6 CO2 pump; 7 back pressure regulator;
8 pre-expansion vessel; 9 micrometric valve with a heating system; 10 temperature
controller; 11 nozzle; 12 expansion vessel.
The operational conditions pre-expansion pressure and temperature ranges of
10-35 MPa and 313.15-323.15 K were evaluated. Ethanol concentration of 27.1 % (w/w)
was employed in all experiments. Different mass ratios between core material and
Polietileneglycol (PEG) of 1:4, 1:6 and 1:10 were tested. The amount of CO2 required for
filling the RESS system was 8.43 g. All experiments were done in duplicate.
158
The residual solvent (ethanol) in the particles was measured by the recording of
the weight loss after the heating of a known amount of sample at 363.15 K for 24 hours.
The encapsulation efficiency was verified by a UV–Vis spectrophotometer
(Hitachi, model U-3010, Tokyo, Japan). First, a sample of core material encapsulated into
PEG was weighed. It was assumed that the ratio between core material and encapsulant
material remained constant after the precipitation. Afterwards, the sample was dissolved in
a suitable solvent for both substances (core material and PEG) and the absorbance was
measured at maximum absorbance wavelength. For the construction of the standard curves
of absorbance vs. concentration in the solvent, hydrochloric acid/potassium chloride buffer
pH 1.0 was used for the determination of anthocyanin encapsulation efficiency.
Comparing the results with a standard curve of absorbance vs. concentration of
anthocyanin encapsulation in the solvent hydrochloric acid/potassium chloride buffer pH
1.0, the Percentage of Encapsulated [PE (%)] and Encapsulation Efficiency [EE (%)] in
each assay were evaluated by the following expressions:
PE (%) =
mass of core material encapsulated
X 100
mass of encapsulated extract system (core + encapsulant materials )
EE (%) =
mass of core material encapsulated
X 100
theoretical mass of core material encapsulated
(1)
(2)
The characteristics of dispersion of the anthocyanin extract in the polymeric
encapsulant material PEG was analyzed by color difference between them using optical
micrographs (Nikon, mod Eclipse E200, Tokyo, Japan).
8.2.3.1.1 Evaluation of the antioxidant activity of the encapsulated anthocyanin extract
after RESS process
159
In order to verify if the evaluated RESS parameters (pressure and temperature)
may affect the anthocyanin extract biological activity, the antioxidant activities of
encapsulated anthocyanin extracts and the anthocyanin extract were compared.
The evaluation of antioxidant activity of these systems was based on the
coupled oxidation of β-carotene and linoleic acid. The technique developed by Marco16
consisted of measuring the bleaching of β-carotene resulting from oxidation by the
degradation products of linoleic acid. One milligram of β-carotene (97 %, Sigma-Aldrich,
St. Louis, USA) was dissolved in 10 cm3 of chloroform (99 %, Ecibra, Santo Amaro,
Brazil). The absorbance was tested after adding 0.2 cm3 of the solution to 5 cm3 of
chloroform, then measuring the absorbance of this solution at 470 nm using a UV–Vis
spectrophotometer (Hitachi, model U-3010, Tokyo, Japan). A measurement between 0.6
and 0.9 indicated a workable concentration of β-carotene. One cm3 of β-carotene
chloroform solution was added to a flask that contained 20 mg of linoleic acid (99%,
Sigma-Aldrich, St. Louis, USA) and 200 mg Tween 40 (99%, Sigma-Aldrich, St. Louis,
USA). Chloroform was removed using a rotary evaporator (Laborota, model 4001,
Vertrieb, Germany), with vacuum control (Heidolph Instruments Gmbh, Vertrieb,
Germany) and a thermostatic bath at 40 °C; then 50 cm3 of oxygenated distilled water
(oxygenation for 30 minutes) was added to the flask with vigorous agitation to form an
emulsion. Five cm3 of the emulsion was added to 0.2 cm3 of the antioxidant solution
(approximately 7.5 mg of free or encapsulated extract; or pure synthetic BHT/1 cm3 of
distilled water) in assay tubes. To the control solution, 0.2 cm3 of pure distilled water was
added. A blank consisting of 20 mg of linoleic acid, 200 mg of Tween 40 and 50 cm3 of
oxygenated distilled water was used to bring the spectrophotometer to zero. Tubes were
manually shaken, and absorbance measurements made at 470 nm immediately after the
addition of the emulsion to the antioxidant solution. The tubes were placed in a water bath
(model TE 159, Tecnal, Piracicaba, Brazil) at 50 ºC. Absorbance measurements were made
at 30 minutes intervals during 2 hours. The average deviation of duplicated experiments
never exceeded 8%, therefore, no additional statistical analysis was considered necessary.
160
8.2.3.2 By conventional method
The conventional method for encapsulation used in this study was the ionic
gelification. The anthocyanin extract was encapsulated by entrapment in Ca-alginate beads
(2.7 mm average diameter). Ca-alginate beads containing anthocyanin extract were
prepared by dripping an adequate volume of a solution 1.5 % of sodium alginate (CD1125,
Vetec Ltda, Rio de Janeiro, Brazil) with an anthocyanin extract concentration of 20 mg/L in
a 20 g/L CaCl2 (Cod 11692, Nuclear Ltda, Brazil), using a 19-G needle (1.5 inch) and a 20
mL syringe (Figure 8.2.3.2.1). The beads were left for 30 min in the CaCl2 solution, then
the beads were separated from the solution and washed with distilled water.
For encapsulation efficiency determination into the beads two methods were
employed: 1) a method based on the complete dissolution of the Ca-alginate beads in
phosphate buffer solution pH 7.417 or sodium citrate solution18; 2) an alternative method
based on the concentration of non encapsulated anthocyanin extract. Samples of the crosslinking solution were taken after encapsulation process. The absorbance of this solution
measured by a UV spectrophotometer (Hitachi, model U-3010, Tokyo, Japan) was
compared with the absorbance of solutions of known concentration of CaCl2 and
anthocyanin extract at maximum absorbance wavelength. A calibration curve was
constructed to correlate the measured absorbance with the concentration of non
encapsulated anthocyanin extract. The Encapsulation Efficiency [EE (%)] was determined
by Equation 3:
EE (%) =
mg anthocyanin extract added / L − mg non encapsulated anthocyanin extract / L
(3)
mg anthocyanin extract added / L
161
Figure 8.2.3.2.1 - Encapsulation by entrapment in Ca-alginate beads.
8.2.4 Anthocyanin Stabilization Studies
8.2.4.1 Free and encapsulated extract degradation studies
After being accurately weighed, encapsulated and free extracts were placed in 3
different environments: ambient temperature (around 25 º C) with or without light, and at 4
ºC without light. At specific time samples were collected and their absorbances were
measured at maximum absorbance wavelength, using distilled water as a blank at a UV
spectrophotometer (Hitachi, model U-3010, Tokyo, Japan).
The sample preparation for the free extracts consisted in dissolving a known
weight of free extract in water in a proportion of 2 mg of extract for 1 mL of distilled water.
Two dilutions of the sample were prepared: one with hydrochloric acid/potassium chloride
buffer pH = 1.0 and the other with sodium acetate/acetic acid buffer pH = 4.5. The pH
values of the buffers were measured using a pH-meter (Digimed, model DM-22, São Paulo,
Brazil) calibrated with buffers at pH 4.01 and 6.86 and they were adjusted with HCl (99.5
162
% Ecibra, Santo Amaro, Brazil). Aliquots of extract were brought to pH 1.0 and 4.5; 15
min later, the absorbance of each equilibrated solution was measured at the maximum
absorption wavelength and 700 nm for haze correction using a 1 cm path length glass cells
(l). The dilution factor (DF) was determined (final volume per original sample volume).
The difference in absorbance values (A) at pH 1.0 and 4.5 (Equation 4) is directly
proportional to the anthocyanin concentration19.
A = (A max − A 700 )pH1,0 − (A max − A 700 )pH 4,5
(4)
In Equation 4, Amax is the absorbance at 512 nm (for jabuticaba skin extract),
and A700 is the absorbance at 700 nm.
For the encapsulated systems a known weight of encapsulated extract was
added in hydrochloric acid/potassium chloride buffer (pH = 1.4, stomach pH) in a heated
bath at 37 ºC and left for 1.5 h. The dissolved encapsulant materials, when necessary, were
separated and the supernatant solution was analyzed by a UV spectrophotometer (Hitachi,
model U-3010, Tokyo, Japan) at maximum absorbance wavelength (512 nm) and 700 nm,
using distilled water as a blank. The amount of released anthocyanins was analyzed using a
calibration curve.
In order to investigate the degradation mechanism of free anthocyanin extract a
first-order reaction kinetic model was applied. The first-order reaction rate constants (k)
and half-lives (t1/2) were calculated according to the following equations:
⎛C ⎞
ln⎜⎜ t ⎟⎟ = − kt + C1
⎝ C0 ⎠
t1/ 2 =
ln 2
k
163
(5)
(4)
where C0 is the initial anthocyanin content and Ct is the anthocyanin content at
the reaction time t.
8.2.4.2 Thermal analysis of free and encapsulated extracts
Free anthocyanin extract, Ca-alginate beads, PEG, anthocyanin extract
encapsulated in Ca-alginate beads and anthocyanin extract encapsulated in PEG were
studied by Differential Scanning Calorimetry (DSC) in order to study the stability of the
extract after encapsulation.
DSC studies were performed using a Shimadzu Differential Scanning
Calorimetry (DSC-50, Shimadzu, Tokyo, Japan). The samples were scanned in sealed
aluminium pans. DSC thermograms were scanned in the first heating run at a constant rate
of 10 ºC/min and a temperature range of 30-500 ºC.
8.2.5 Anthocyanin Extract Release Studies
Similar procedure to the described in section 8.2.4.1 was done for anthocyanin
extract release studies, differing that periodically the supernatant solution was analyzed.
The average deviation of triplicate experiments never exceeded 8%, therefore, no additional
statistical analysis was considered necessary.
8.2.6 Statistical Analysis
For establishing the statistical significant differences or similarities between the
values of Percentage of Encapsulated [PE (%)], the analysis of variance (Tukey test) was
used. A confidence coefficient of 95 % was used for the comparison of all the mean’s pairs.
164
8.3 Results and discussion
8.3.1 Encapsulation of anthocyanin extract by RESS process
The color of PEG (transparent gray - Figure 8.3.1.1a) is obviously different
from that of anthocyanin extract (red), so we analyzed the characteristics of dispersion of
anthocyanin in the encapsulant material by color difference between anthocyanin extract
and PEG using optical micrographs.
The optical micrographs (Figure 8.3.1.1b and 8.3.1.1c) display the dispersion
characteristics of anthocyanin extract in the anthocyanin extract/PEG microparticles
obtained by the RESS process.
Generally most polymers could absorb a large concentration of CO2 (about 10–
40 wt %) that either swells the polymers or melts them at a temperature much below theirs
melting/glass transition temperature (about 10–50 K)20. PEG used in this work has low
melting/glass transition temperature (333–335 K), so its precipitates obtained at higher
temperature (323.15 K) tend to be soft, sticky and easily agglomerated resulting in
formation of bigger coalesced particles, as showed in Figure 8.3.1.1c, than that obtained at
313.15 K (Figure 8.3.1.1b) and with a higher anthocyanin extract loading (Table 8.3.1.1).
165
Figure 8.3.1.1 - Optical micrographs of unprocessed PEG (a), Encapsulated anthocyanin
extract – T: 313.15 K; P: 100 bar (b), Encapsulated anthocyanin extract – T: 323.15 K; P:
100 bar (c)
166
In order to explore the effect of temperature and pressure on the anthocyanin
extract loading (Percentage of Encapsulated [PE (%)]) we performed experiments in the
range of 313.15-323.15 K and 10-35 MPa. With the increase of temperature, PE increases.
With the increase of temperature the precipitates of PEG tend to be softened due to the
effect of supercritical CO2 on it, i.e. it becomes difficult for PEG to be hardened in the rapid
expansion of the supercritical solution (PEG + ethanol + anthocyanin extract + supercritical
CO2) resulting in the increase of anthocyanin extract loading in the precipitated particles.
Indeed, with the increase of pressure, PE increases, probably due to the increase of CO2
density. Nevertheless, it can be observed that the effect of temperature was more
pronounced than that of the pressure: increasing the temperature from 313.15 to 323.15 K
promoted an increase in PE greater than that provoked by the increase in pressure.
On the other hand, after expansion the produced anthocyanin extract/PEG
particles did not tend to agglomerate as expected because the residual co-solvent (ethanol)
acts as a good nonsolvent for the polymer. The amount of the residual ethanol in the
microcapsules was 0.40 % (w/w). Matsuyama et al.21 obtained an amount of residual
ethanol in their microcapsules less than 1 % (w/w) for different polymers.
The Tukey test for data in Table 8.3.1.1 demonstrated that for the PE obtained
at 323.15 K, the differences were not statistically significant (5%) while for the PE obtained
at 313.15 K, the differences were statistically significant only at 10 MPa. Based on this, we
can choose as best operating RESS process conditions for anthocyanin extract
encapsulation 313.15 K and 20 MPa.
Table 8.3.1.1 - Percentage of Encapsulated [PE (%)] of encapsulated anthocyanin extract
obtained by RESS process
T (K)
PE (%)
10
20
313.15
23.86
24.69
24.98
25.17
323.15
24.70
24.87
25.36
25.94
167
30
35 (MPa)
The concentration of PEG added was found to be an important factor that
influences the Percentage of Encapsulated in the PEG microparticles. At 313.15 K and 20
MPa the RESS process produced different PE and Encapsulation Eficiency [EE (%)] when
different proportions of anthocyanin extract:PEG were employed. Figure 8.3.1.2 shows that
a reduction in PEG concentration enhances both values (PE and EE), reaching the highest
values, 27.65 % and 79.78 %, respectively.
Figure 8.3.1.2 - Influence of different proportions of anthocyanin extract:PEG on the
percentage of encapsulated (gray bars); encapsulation efficiency (black bars)
Anthocyanin extract encapsulated by RESS retained its dark red color,
indicating no degradation of the dye during the encapsulation procedure. Since pigment
degradation is, according to Mattea et al.22, directly related to color and biological activity
loss, the evaluation of the antoxidant activities of encapsulated anthocyanin extracts and the
anthocyanin extract were compared. Figure 8.3.1.3 confirms that no degradation, i.e. lost of
antioxidant activity has occurred. It can also be noted that recognized pure synthetic BHT
presented higher antioxidant activities than anthocyanin extract.
168
Figure 8.3.1.3 - Antioxidant activity of encapsulated anthocyanin extracts obtained using
different operating RESS conditions (gray symbols), the anthocyanin extract (♦) and pure
synthetic BHT (■); without any antioxidant compound (▲)
Corroborating our findings, Jacobson et al.15 also demonstrated that particle
formation processes using supercritical CO2 can retain the antioxidative activity of the
compound or class of compounds of interest after processing.
8.3.2 Encapsulation of anthocyanin extract by conventional method
It was not possible to quantify the encapsulation efficiency using the method
based on the complete dissolution of the Ca-alginate beads in phosphate buffer solution pH
7.417 or sodium citrate solution18. In both cases a change of anthocyanin extract color from
red to brown was observed indicating anthocyanin degradation. Thus, an alternative method
based on the concentration of non encapsulated anthocyanin extract was used.
169
The loss of anthocyanin extract during the encapsulation process was less than
1.33 %, so the Encapsulation Efficiency [EE (%)] was 98.67 %. Most physical
encapsulation technologies can give a loading capacity as high as 99 %. Numerous articles
report Ca-alginate as a successful carrier for different compounds in its wet or dry form23,24
and usually it demonstrates a high encapsulation efficiency3.
8.3.3 Anthocyanin stabilization by encapsulation
The free and encapsulated extract degradation studies demonstrated that
different environments as light and temperature interfere in the anthocyanin stability, as
expected (Figures 8.3.3.1 and 8.3.3.2).
Figure 8.3.3.1 - Degradation of free anthocyanin extract at different environments.
170
The free anthocyanin extract presented a first order decomposition curve for all
the samples (Table 8.3.3.1). First order kinetics for degradation of anthocyanins has been
reported on different sources as black raspberry, sour cherry, concord grape, red cabbage,
radish and strawberry25. Although, the anthocyanin degradation time differs depending on
conditions as pH, storage temperature, anthocyanin chemical structure and concentration,
light, oxygen and others26.
Table 8.3.3.1 - Kinetic parameters of free anthocyanin extract degradation obtained by the
first order kinetic model and half-life degradation equation
k
-1
(days )
C1
R2
t(1/2)
(days)
Low temperature
0.0442
-0.0115
0.947
15.68
Without Light
0.0440
0.0194
0.914
15.75
Low Temperature
0.0618
0.0083
0.877
11.22
Figure 8.3.3.2 - Degradation of encapsulated anthocyanin extract at different environments
by RESS process and conventional method.
171
Both encapsulated systems were less influenced by the environment than the
free extract (Figure 8.3.3.2). The degradation of the free extract occurred faster than the
encapsulated, possibly due the protection of the anthocyanin molecules when encapsulated
by the polymeric matrices preventing oxidation.
After 14 days the encapsulated extract by conventional method into Ca-alginate
beads stored at low temperature (4 ºC) showed no significant degradation, and the one kept
at ambient temperature and light presented around 20% and 50% of degradation rate,
respectively. Meanwhile, the encapsulated extract by RESS process into PEG showed
degradation rate values a little higher. To better understand these results, DSC analyses of
both encapsulated systems, of both encapsulant materials and the free anthocyanin extract
were done.
The DSC analysis (Figure 8.3.4.3) shows different endothermic peaks for the
encapsulated anthocyanin extract/PEG system, which seems to be related to the
anthocyanin extract and PEG, indicating that probably this encapsulated system is a not
good mixture of both compounds. In contrast, only one endothermic and different peak
from the Ca-alginate beads and the anthocyanin extract was observed for the encapsulated
anthocyanin extract/Ca-alginate system, indicating that polymer and extract are well
mixed27. This also suggests that a chemical interaction between the Ca-alginate beads and
the extract can occur instead of a simple physical mixture. Given that the anthocyanin
extract/Ca-alginate peak at a higher temperature may confirm the better stability of this
encapsulated system under different environments.
172
Figure 8.3.4.3 - DSC thermograms of free anthocyanin extract, Ca-alginate beads, PEG,
anthocyanin extract encapsulated in Ca-alginate beads and anthocyanin extract
encapsulated in PEG.
8.3.4 Release studies
The release of anthocyanin was studied in hydrochloric acid/potassium chloride
buffer solution of pH 1.4. Figure 8.3.4.1 shows that colorant is released faster in the first
173
hour and even after 150 min the beads still had the characteristic color of the anthocyanins
for the encapsulated particles obtained by conventional method. On the other hand, for the
encapsulated particles obtained by RESS process the complete release occur in the first 15
minutes.
Figure 8.3.4.1 - The cumulative release of anthocyanin extract from the encapsulated
systems obtained by RESS process and conventional method at pH 1.4 and temperature 37
ºC.
Sequential release study was performed for the encapsulated particles obtained
by conventional method in order to evaluate the influence of the medium saturation in the
release of colorant. The higher release occurred at 30 min decreasing until values next to 0
in the period of 2 hours. After that time the beads were still colored but very small release
was detected. Similar results were achieved by different authors when studying the release
from pure Ca-alginate beads and Ca-alginate beads with small percentage of chitosan in its
composition17,28,29. Only around 5 – 20% of the bead content was released in low pH
medium at 37 ºC, it was influenced by the chemical composition of the encapsulated
compound and the amount of chitosan presented in the Ca-alginate beads. These results
174
probably are associated to the capacity of the Ca-alginate beads to promote controlled
release, thus the slower and not complete release of anthocyanin extract from this
encapsulant material at the evaluated conditions can be interesting for pharmaceutical and
cosmetic purposes.
8.4 Conclusions
The encapsulation of anthocyanin extract obtained from jabuticaba skins, a
Brazilian potential source of anthocyanins, in PEG by RESS process and Ca-alginate
matrix by conventional method were successfully accomplished. The encapsulation
efficiency of the extract in Ca-alginate beads was higher (98.67 %) than those obtained by
RESS process (79.78 %). Encapsulated particles made by RESS at different pressure and
temperature conditions have been tested with this assay, all showing retained biological
activity. Pressure, temperature and encapsulant material concentration were operating
parametes that affected the RESS process, being this process influenced mainly by
temperature. The best operating RESS process condition for anthocyanin extract
encapsulation was determinated at 313.15 K and 200 bar. The degradation studies indicated
that both encapsulated systems were more stable to the light and temperature than the free
extract, and the encapsulated anthocyanin extract obtained by conventional method was
more stable to that obtained using supercritical CO2, probably due to a chemical interaction
between the extract and the encapsulant material Ca-alginate. The release of the
encapsulated anthocyanin extract in Ca-alginate in acid buffer solution was incomplete and
slower than in PEG, indicating that Ca-alginate beads can promote controlled release,
which is interesting for pharmaceutical and cosmetic purposes.
175
Acknowledgements
The authors are grateful to CNPq for the financial support and to the University of
Campinas.
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179
180
CAPÍTULO 9 - CONCLUSÕES GERAIS
Os resultados apresentados nesta tese demonstraram que a unidade construída
possibilita a extração, micronização e estabilização de pigmentos funcionais por
encapsulação em matrizes poliméricas utilizando fluidos pressurizados. Comparando os
resultados obtidos utilizando a unidade construída com os obtidos utilizando equipamentos
comerciais e equipamentos construídos por outros grupos de pesquisa foi verificado que a
unidade construída produz resultados confiáveis e reprodutíveis. Os processos modelos
para as etapas de validação escolhidas foram: i) para o processo de extração – extração de
pigmentos de sementes de urucum utilizando CO2 supercrítico puro; ii) para o processo de
formação de partículas – a formação de partículas não encapsuladas dos pigmentos
quercetina e β-caroteno.
A viabilidade técnica de se desenvolver processos de extração na unidade
construída utilizando-se das técnicas: i) extração assistida por ultrassom; ii) extração
fracionada; iii) extração com líquidos pressurizados; iv) e extração assistida por CO2 a alta
pressão, foram avaliadas utilizando como fontes vegetais pigmentares cascas de jabuticaba
e sementes de urucum. Dentre os quatro processos avaliados somente o processo de
extração assistida por ultrassom não foi desenvolvido eficazmente. De acordo com os
resultados inferiu-se que a potência ultrassônica do banho de ultrassom utilizado não foi
suficientemente alta para poder causar os benefícios que a irradiação ultrassônica propicia
no processo extrativo. Portanto, foi sugerido que possivelmente, com pequenas
modificações, tais como o aumento da potência das ondas ultrassônicas através da
incorporação de mais transdutores, uma vez que a parede da célula extratora reduz a
permeação da radiação; ou a realocação dos transdutores visando à redução da distância
entre os transdutores e a matéria-prima os processos de extração que utilizam solventes
pressurizados assistidos com ultrassom poderiam ser desenvolvidos eficientemente.
Uma vez que os outros processos de extração foram possíveis de serem
desenvolvidos na unidade construída, a potencialidade dos outros três processos aplicada à
181
extração de pigmentos funcionais sem a sua degradação durante o processo extrativo foi
explorada.
A extração fracionada de cascas de jabuticaba produziu dois extratos com
atividades antioxidantes: um rico em pigmentos antociânicos (utilizando etanol
pressurizado) e outro rico em compostos lipofílicos que podem ser compostos da classe dos
flavonóides ou dos óleos essenciais (utilizando CO2 supercrítico puro).
O rendimento da extração com etanol pressurizado de cascas de jabuticaba na
condição otimizada (Pressão de 50 bar, Temperatura de 80 ºC e Tempo de extração estática
de 9 minutos) foi significativamente maior do que quando utilizando o método de extração
convencional, percolação em leito fixo, utilizando o mesmo solvente sob pressão e
temperatura ambientes. As variáveis resposta: rendimento de extração, extração de
antocianinas e compostos fenólicos totais foram afetadas positivamente pelo aumento de
temperatura, porém o uso de temperaturas maiores que 80 ºC por um maior período de
tempo (maior que 9 minutos de extração estática) causou degradação das antocianinas
durante o processo extrativo.
O rendimento da extração assistida com CO2 a alta pressão na condição
otimizada (Pressão de 117 bar, Temperatura de 80 ºC e Relação Volume de CO2/Volume de
Solvente+Matéria-Prima de 20 %) foi significativamente maior do que quando utilizando o
método de extração com líquido pressurizado com o mesmo solvente (água acidificada em
pH 2,5) sob a uma mesma pressão e temperatura. Todos os parâmetros de processo
afetaram significativamente a extração de compostos fenólicos totais, enquanto para a
extração de antocianinas somente pressão e temperatura tiveram efeitos significativos.
Comparando-se os resultados de extração de antocianinas e outros compostos
fenólicos de cascas de jabuticaba utilizando o processo de extração com etanol pressurizado
com os obtidos utilizando o processo de extração assistida com CO2 a alta pressão, ambos
em condições otimizadas, conclui-se que este último é capaz de extrair mais eficientemente
estes tipos de compostos.
A viabilidade técnica de a unidade construída desenvolver processos de
formação de partículas encapsuladas: i) de extrato de sementes de urucum ricos no
182
pigmento funcional bixina; ii) de extrato de cascas de jabuticaba ricos em pigmentos
antociânicos; iii) e de pigmento rutina em sua forma pura, empregando CO2 supercrítico
como solvente (via RESS) ou anti-solvente (via SAS) foram avaliadas utilizando como
material de encapsulação o polímero polietilenoglicol (PEG). Os 3 processos avaliados
foram desenvolvidos eficazmente, entretanto, a potencialidade do processo de encapsulação
utilizando fluidos pressurizados proporcionar uma maior estabilidade a estes pigmentos
instáveis, bem como não os degradar durante o processo de encapsulação foi explorada
somente para o caso das antocianinas.
Extrato rico em antocianinas obtido através do emprego de etanol pressurizado
utilizando a unidade construída foi estabilizado através da sua encapsulação no polímero
PEG empregando CO2 supercrítico como solvente (via RESS) e etanol como co-solvente
visando aumentar a solubilidade do material de encapsulação no fluido supercrítico. O
estudo de degradação indicou que quando encapsulado pelo método RESS o extrato
antociânico foi mais estável do que quando não encapsulado e menos estável do que
quando encapsulado pelo método convencional de gelificação iônica em alginato de cálcio,
possivelmente devido à interação química que este último método propiciou entre o extrato
e a matriz polimérica. Adicionalmente, também se verificou que o desenvolvimento do
processo de formação de partículas encapsuladas via RESS não afeta a estabilidade do
extrato antociânico, mantendo, portanto, as suas atividades biológicas.
183
184
SUGESTÕES PARA TRABALHOS FUTUROS
i)
Avaliar economicamente os processos de extração e formação de partículas
desenvolvidos neste trabalho na unidade construída utilizando simuladores
comerciais (SuperPro Designer®, etc.);
ii)
Otimizar o processo de encapsulação/co-precipitação de extrato rico em
carotenóides
oriundo
de
sementes
de
urucum,
com o
polímero
polietilenoglicol (PEG) via SAS utilizando a unidade construída e CO2
supercrítico como anti-solvente comparando os resultados relacionados à
estabilidade com os obtidos por outros métodos;
iii)
Avaliar a incorporação de um sistema que possibilite um desenvolvimento
eficiente de extrações com fluidos pressurizados assistidas com ultrasom na
unidade construída;
iv)
Verificar a viabilidade técnica da utilização da unidade construída na
secagem de extratos aquosos pigmentares ou não;
v)
Verificar a viabilidade técnica da extração com fluidos pressurizados e
formação de partículas bioativas não encapsuladas em uma única etapa (“inline”) utilizando a unidade construída;
vi)
Desenvolver novos dispositivos/atomizadores que melhorem a formação de
partículas utilizando fluidos supercríticos como solventes ou anti-solventes
na unidade construída.
185
186
MEMÓRIA DO PERÍODO DO DOUTORADO
Diego Tresinari dos Santos vem desenvolvendo o presente projeto de pesquisa, com bolsa e
auxílio financeiro do CNPq (Edital CNPq 070/2008 e Edital MCT/CNPq 15/2007 Universal - Faixa B, respectivamente). Até o presente momento 23 créditos foram cursados,
sendo 17 créditos referentes a disciplinas (Termodinâmica, Fenômenos de Transporte I,
Fenômenos de Transporte II, Tópicos Especiais em Físico-Química X e Seminários) e 6
créditos referentes a 3 participações no Programa de Estágio Docente grupo C - PED C
(segunda participação com bolsa e com carga didática de 20 %), sendo obtido um
Coeficiente de Rendimento (CR) de 3,7391. Os procedimentos experimentais do projeto de
pesquisa se iniciaram após participação em treinamentos organizados pelos integrantes com
mais experiência no laboratório. O aluno também participou em treinamentos referentes à
utilização de softwares para simulação de processos, tais como Tecanalysis e SuperPro
Designer® 6.0. Durante seu trabalho acadêmico Diego participou de 4 congressos
científicos (CBPOL 2009, ENEMP 2009, SLACA 2009 e PROSCIBA 2010), tendo
apresentado 2 trabalhos na forma de painéis e 1 oralmente. Interagiu durante o período do
doutorado com pesquisadores: i) da Escola de Engenharia de Lorena da USP no
desenvolvimento do projeto intitulado Avaliação da Atividade Antimicrobiana de Produtos
Naturais para Obtenção de Novos Biofármacos: Fase 1 - Estudo dos extratos brutos e sua
associação financiado pela FAPESP; ii) da Faculdade de Engenharia Química da
UNICAMP no estudo da encapsulação e estabilidade das antocianinas, bem como na
avaliação do reaproveitamento do resíduo da extração supercrítica cascas de banana para a
remoção de metais pesados; iii) do Centro Pluridisciplinar de Pesquisas Químicas,
Biológicas e Agrícolas (CPQBA) da UNICAMP no estudo de identificação dos tipos de
antocianinas presentes nos extratos de jabuticaba e jambolão por espectrometria de massas,
bem como na avaliação da hidrólise de saponinas glicosiladas por água subcrítica + CO2
visando uma melhor identificação destes compostos por espectrometria de massas; iv) da
Faculdade de Ciências Farmacêuticas da USP no estudo da micronização de fármacos não
encapsulados e encapsulados em ciclodextrinas via RESS e SAS utilizando a unidade
187
multipropósito construída; v) bem como com seus colegas de laboratório em diferentes
trabalhos de pesquisa envolvendo a extração de compostos bioativos de fontes vegetais.
188
PRODUÇÃO BIBLIOGRÁFICA
Artigos completos publicados/aceitos em periódicos
1) CAVALCANTI, R. N. ; SANTOS, D. T. ; MEIRELES, M. A. A. . Non-thermal
Stabilization Mecanisms of Anthocyanins in Model and Food Systems - an overview. Food
Research International, In press, 2011.
2) SANTOS, D. T. ; MEIRELES, M. A. A. . Extraction of Volatile Oils by Supercritical
Fluid Extraction: Patent Survey. Recent patents in engineering, In press, 2011.
3) SANTOS, D. T. ; VEGGI, P. C. ; MEIRELES, M. A. A. . Extraction of Antioxidant
Compounds from Jabuticaba (Myrciaria cauliflora) Skins: Yield, Composition and
Economical Evaluation. Journal of Food Engineering, v. 101, p. 23-31, 2010.
4) SANTOS, D. T. ; MEIRELES, M. A. A. . Carotenoid Pigments Encapsulation:
Fundamentals, Techniques and Recent Trends. The Open Chemical Engineering Journal, v.
4, p. 42-50, 2010.
5) SANTOS, D. T. ; MEIRELES, M. A. A. . Jabuticaba as a Source of Functional
Pigments. Pharmacognosy Reviews, v. 3, p. 127-132, 2009.
Trabalhos completos publicados em anais de congressos
1) NAVARRO-DÍAZ, H. J. ; SANTOS, D. T. ; MEIRELES, M. A. A. . Phenolic
Compounds Recovery From Punica granatum L. Leaves Using Supercritical CO2. In: II
Iberoamerican Conference on Supercritical Fluids (PROSCIBA), 2010, Natal-RN. Full
Paper, 2010. p. 1-7.
189
2) VEGGI, P. C. ; SANTOS, D. T. ; MEIRELES, M. A. A. . Extraction of antioxidant from
some Brazilian plants. In: 12th European Meeting on Supercritical Fluids, 2010, GrazAustria. Proceedings of the 12th European Meeting on Supercritical Fluids, 2010.
3) ALBUQUERQUE, C. L. C. ; SANTOS, D. T. ; MEIRELES, M. A. A. . Functional
properties of supercritical extract from ylang-ylang (Cananga odorata) peel fruit. In: 12th
European Meeting on Supercritical Fluids, 2010, Gras-Austria. Proceedings of the 12th
European Meeting on Supercritical Fluids, 2010.
4) ALBARELLI, J. Q. ; MACUMOTO, A. ; CARVALHO, L. ; SANTOS, D. T. ;
MEIRELES, M. A. A. ; BEPPU, M. M. . Encapsulação de Corantes Funcionais em Matriz
de Alginato Puro ou Recoberto por Biopolímeros. In: 10º Congresso Brasileiro de
Polímeros, 2009, Foz do Iguaçú-PR. Anais do 10º Congresso Brasileiro de Polímeros,
2009. p. 1-6.
Resumos publicados/aceitos em anais de congressos
1) SOUZA, I. P. S. ; CARVALHO, G. H. ; SANTOS, D. T. ; MEIRELES, M. A. A. .
Estudo da influência do método de extração na obtenção de extratos de semente de
Myrciaria cauliflora. In: XVIII Congresso Interno de Iniciação Científica da UNICAMP,
20010, Campinas-SP. Caderno de Resumos, 2010. p. 136-136.
2) FERREIRA, J. A. ; CARVALHO, G. H. ; SANTOS, D. T. ; MEIRELES, M. A. A. .
Caracterização química dos extratos de semente de jabuticaba obtidos por diferentes
tecnologias. In: XVIII Congresso Interno de Iniciação Científica da UNICAMP, 20010,
Campinas-SP. Caderno de Resumos, 2010. p 137-137.
190
3) SANTOS, D. T. ; VEGGI, P. C. ; MEIRELES, M. A. A. . Obtaining anthocyanin-rich
extracts from jabuticaba skins: technical and economical evaluation. In: The 15th World
Congress of Food Science and Technology (IUFOST), 2010, Cape Town-Africa do Sul.
Proceedings of the 15th World Congress of Food Science and Technology, 2010.
4) SANTOS, D. T. ; VEGGI, P. C. ; MEIRELES, M. A. A. . Avaliação Técnico-Econômica
da Extração de Antocianinas por Percolação em Leito Fixo. In: XXXIV Congresso
Brasileiro de Sistemas Particulados (ENEMP), 2009, Campinas-SP. Livro de Resumos,
2009. p. 306-306.
5) SOUZA, J. S. B. ; MALVEZZI, C. K. ; FREITAS, J. E. ; MEIRELES, M. A. A. ;
SANTOS, D. T. ; SILVA, S. S. . Avaliação Comparativa da Atividade Antimicrobiana do
extrato supercrítico das folhas de Psidium guajava frente à Staphylococcus aureus e
Escherichia coli. In: I Semana de Biotecnologia Industrial - Inovações e Perspectivas da
Biotecnologia Industrial no Brasil, 2009, Lorena-SP. Livro de Resumos, 2009. p. 1-1.
6) VAZ, N. ; SANTOS, D. T. ; MEIRELES, M. A. A. . Composição Química do Extrato de
Folhas de Jaca (Artocarpus heterophyllus). In: 8º Simpósio Latino Americano de Ciência de
Alimentos (SLACA), 2009, Campinas-SP. Cd..., 2009. p. 1-1.
7) CAVALCANTI, R. N. ; SANTOS, D. T. ; MEIRELES, M. A. A. . Composição
Centesimal das Partes Comestíveis do Fruto de Syzygium cumini. In: 8º Simpósio Latino
Americano de Ciência de Alimentos (SLACA), 2009, Campinas-SP. Cd..., 2009. p. 1-1.
191
192
APÊNDICE I - ARTIGO DE REVISÃO - JABUTICABA AS A SOURCE OF
FUNCTIONAL PIGMENTS
Publicado no periódico Pharmacognosy Reviews.
193
194
Phcog Rev. Vol, 3, Issue 5, 127-132, 2009
Available Online : www.phcogrev.com
PHCOG REV. : Review Article
Jabuticaba as a Source of Functional Pigments
Diego T. Santos and M. Angela A. Meireles*
Laboratory of Supercritical Technology: Extraction, Fractionation, and Identification of vegetable extracts (LASEFI),
Department of Food Engineering (DEA), University of Campinas (UNICAMP), Rua Monteiro Lobato, 80, Cid. Universitária
Zeferino Vaz, 13083-862, Campinas - SP, Brazil
Author for correspondence*: [email protected]; Phone: 00551935214033
ABSTRACT
The relatively high concentration of anthocyanins in the diet is of prospective importance to human health. Anthocyanins
contribute greatly to the antioxidant properties of certain colorful foods, such as grapes and cranberries. Many studies in recent
years have focused on the study of these functional pigments from different perspectives. The present review highlights recent
studies on the health-promoting properties of anthocyanins. It presents latent anthocyanin sources and demonstrates the
potentiality of an under-utilized non-conventional source widespread in Brazil called jabuticaba (Myrciaria cauliflora).
KEY WORDS: Anthocyanins, functional pigments, health-promoting properties, jabuticaba.
INTRODUCTION
Oxidative reactions in the human body have been appointed
as the cause of diseases initiation and progression. The
damage generated by free radicals and reactive oxygen species
has been linked to some neurodegenerative disorders and
cancers, and oxidation of low-density lipoprotein is a major
factor in the promotion of coronary heart disease (CHD) and
atherosclerosis (1).
Epidemiological evidences suggest that a diet high in fruits
and vegetables plays an important hole in reducing the
incidence of many oxidative and inflammatory diseases (2, 3).
The reason of this accomplishment can be the coloration of
these foods, in general, a rich source of many phenolic
antioxidants.
The wish of a healthier diet allied with the increasingly
concern of consumers over the use of synthetic additives in
food has pushed the food industry to search for new sources
of natural pigments (4). Anthocyanins are a type of functional
pigments, responsible for a wide range of colors present in
vegetables, flowers, fruits, and derived products. It is known
that anthocyanin pigments act as strong antioxidants and antiinflammatory,
with
antimutagenic
and
cancer
chemopreventative activities (5). These bioactive properties
have been already demonstrated in “in vitro” and “in vivo”
studies (6) and an increase of publications in this area can be
observed in the recent years.
Grapes and berries are well known for their antioxidant
properties due to the presence of anthocyanins, many studies
were done to extract and evaluate these compounds. The
challenge for obtaining this class of pigments in industrial
scale can be achieved by researching under utilized tropical
fruits. Jabuticaba (Myrciaria caulifora) is a Brazilian fruit that can
be potentially used as anthocyanins source because of their
high content. It is denoted by many different names such as
jaboticaba, guaperu, hipavuru, sabará or ybapuru (7).
The present work aims to highlight recent studies on the
health-promoting properties of anthocyanins. As well as,
present new potential anthocyanin sources obtained from
© Phcog.Net 2009 | www.phcog.net
non-conventional plants, giving special attention to a
widespread fruit in Brazil called jabuticaba (Myrciaria cauliflora).
ANTHOCYANINS CHEMISTRY
Phenolic compounds are part of the secondary metabolism of
plants and are of great importance for their survivor in
unfavourable environment. They protect the species against
adverse factors such as drought, UV radiation, infections or
physical damage and regulate their development (8). A class of
phenolic compounds easily found in the Plant Kingdom is the
anthocyanins. They are water-soluble pigments that confer the
bright red, blue, and purple colors of fruits and vegetables
such as berries, grapes, apples, purple cabbage, etc (9).
The basic structures of anthocyanins are the anthocyanidins
(Figure 1). These structures, also known as aglycons, consist
of an aromatic ring [A] bonded to an heterocyclic ring [C] that
contains oxygen, which is also bonded by a carbon–carbon
bond to a third aromatic ring [B] (10). When the
anthocyanidins are found in their glycosylated form (bonded
to a sugar moiety) via the C3 hydroxyl group in ring C they are
known as anthocyanins.
In this way, a huge variety of anthocyanins can be observed
spread in nature only varying in the basic anthocyanidin
skeleton, the position and extent to which the glycosides are
attached to the skeleton. The six most common anthocyanidin
skeletons are cyanidin (Cy), delphinidin (Dp), pelargonidin
(Pg), malvidin (Mv), petunidin (Pt), and peonidin (Pn) (Table
1) (when R1, R2, R3 and R5 are OH). Their distribution in
fruits and vegetables is, respectively: 50%, 12%, 12%, 7%, 7%
and 12% (11).
The most common anthocyanins in nature are the glycosylated
derivatives of the three non-methylated anthocyanidins (Cy,
Dp and Pg). They are found in 80% of pigmented leaves, 69%
in fruits and 50% in flowers being the most common
anthocyanin the Cy-3-glucoside (5).
ANTHOCYANINS HEALTH-PROMOTING
PROPERTIES
Several studies show that a consumption of dietary
phytochemicals, of which anthocyanins form a considerable
127
Diego and Angela, Phcog Rev. 3(1):127-132 [2009]
Figure 1: Structural identification of anthocyanidins (aglycons)
Source
Baguaçu
Bilberry
Blackcurrant
Black Bean
Black Olives
Black Rice
Blackberry
Blueberry
Cherry
Chokeberry
Cranberry
Crowberry
Eggplant
Jambolão
Jabuticaba
Pomegranate (juice)
Port Wine
Purple Corn
Raspberry
Red Cabbage
Red Grape
Red Onion
Red Radish
Rhubarb
Strawberry
Table 1: Sources of anthocyanins
Anthocyanins (mg/100g of fresh weight)
596,4-577,7
214.7-698
128-476
24.1-44.5
42-228
10-493
82.5-325.9
25-495
2-450
311.02-1480
19.8-140
360
8-85
108,8-386
310-315
600-765
14-110
1642
19-687
24,2-322
30-750
23.3-48.5
100-154
4-200
19-55
© Phcog.Net 2009 | www.phcog.net
References
(33)
(34-36)
(37-39)
(40)
(36)
(41)
(42-44)
(34, 35, 45-48)
(49)
(8, 34, 39, 50, 51)
(39, 51, 52)
(51)
(39, 51)
(33, 53)
(53)
(54)
(38)
(55)
(39, 42, 44, 56, 57)
(39)
(29, 58, 59)
(39, 60)
(39, 61)
(38, 51)
(62, 63)
128
Diego and Angela, Phcog Rev. 3(1):127-132 [2009]
Table 2: Articles that have mentioned and/or shown the potential of Jabuticaba as a source of functional pigments published in journals
indexed in the Web of Science and Scopus database
Title
Year
Reference
Carbohydrates, Organic-Acids and Anthocyanins of Myrciaria-Jaboticaba-Berg
1972
(67)
Growth Relations and Pigment Changes in Developing Fruit of Myrciaria Jaboticaba
1996
(70)
Anthocyanin Antioxidants from Edible Fruits
2004
(1)
Application of Tristimulus Colorimetry to Optimize the Extraction of Anthocyanins from Jaboticaba (Myricia
2005
(4)
Jaboticaba Berg.)
Blue Sensitizers for Solar Cells: Natural Dyes from Calafate and Jaboticaba
2006
(71)
Bioactive Depsides and Anthocyanins From Jaboticaba (Myrciaria cauliflora)
2006
(68)
Quantitative Analysis af Antiradical Phenolic Constituents from Fourteen Edible Myrtaceae Fruits
2008
(2)
part, may promote several health benefits. Due to their
injury was induced by the administration of D-galactosamine
biological activity, in particular their antioxidant and antithe anthocyanins also demonstrated this protective effect (21).
inflammatory activities, a reduction in the risk of
Beneficial Effects in Cognitive Performance
cardiovascular disease, diabetes, cancer, an increase of the
Several studies performed in animals have shown that
cognitive performance, and others can be achieved (12-15).
anthocyanins can increase the cognitive performance, and also
protect the brain function by reducing oxidative ischemic
Prevention of Cardiovascular Diseases
Prevention of cardiovascular diseases is possibly the most
damage and enhancing memory (13, 15, 22).
studied effect of anthocyanins in the organism and the one for
Protective Effect on Gastric Damage
which a great quantity of epidemiological evidence exists. This
The protective effect of anthocyanins on gastric damage is
class of phenolic compounds is capable of acting on different
closely related with the capacity of this group of flavonoids to
cells involved in the development of atherosclerosis, one of
prevent or diminish the inflammatory process. It is known
the leading causes to cardiovascular dysfunction (16).
that inflammation implicates, at least initially, in processes of
Anti-diabetes Effects
gastric injury. Studies have shown that cyanidin protects
According to some studies anthocyanins may also prevent
gastric mucosa from the damage caused by aspirin (23).
Cell Regeneration Properties
type 2 diabetes and obesity. They affect the intestinal glucose
absorption by retarding the release of glucose during
Mucopolysaccharides are important to maintain the integrity
of both perivascular tissue and the basal membrane.
digestion, insulin level/secretion/action and lipid metabolism
“in vitro” and “in vivo”. These phytochemicals were found to
Anthocyanins were reported to induce active phagocytosis of
be potent inhibitors of starch digestion, and effective
pigment material and intense cell regeneration in “in vitro”
inhibitors of the a-glucosidase/maltase activity (17).
studies using endothelial cells from human umbilical cord (24).
Anticancer Activity
A growth promoting activity on fibroblasts and on smooth
Although it is not certain that anthocyanins intake reduces
muscle cells was also reported in the same study. The
regeneration of the cellular component of the vessel wall and
cancer risk in humans, the antioxidative capacity of these
functional pigments is well known. Studies suggest that
of the perivascular tissues may be aid by anthocyanins intake,
anthocyanins intake may reduce oxidative damage. A study
due to their stimulating effect on mucopolysaccharides.
performed in Germany showed that individuals who
Beneficial Ocular Effects
consumed an anthocyanins/polyphenolics-rich fruit juice had
Anthocyanins have demonstrated a beneficial impact on the
circulatory system improving the microcirculation of the
reduced oxidative DNA damage and a significant increase in
reduced glutathione when compared to controls (18).
blood and consequently improving vision at dusk and at night.
Anti-inflammatory Effects
Owing to those properties, anthocyanins have been applied in
Beneficial immune responses have been shown in human
the production of ophthalmic preparations for research
endothelial cells upon exposure to anthocyanins. The property
purposes (25).
of reducing the oxidative damage is the manly reason of these
Protective Effect Against Collagen Degradation
Elastase is an important proteolytic enzyme involved in the
results, as inflammation processes are usually accompanied by
degradation of collagen and other components of the
excessive production of reactive oxygen and nitrogen
metabolites. The anti-inflammatory effects could be confirmed
extravascular matrix in certain pathological conditions such as
by analyzing the compound metabolites at doses and
atherosclerosis, pulmonary emphysema, and rheumatoid
comparing to those found in plasma after fruits anthocyaninsarthritis. “In vitro” assays have demonstrated the ability of
rich administration (19).
anthocyanins to inhibit these enzymes acting as a protection
Protective Effect Against Hepatic Damage
against collagen degradation. It is believed that anthocyanins
Anthocyanins have shown to be effective in liver protection
interact with collagen metabolism by cross-linking collagen
fibers and making them more resistant to collagenase action
from hepatotoxicity induced by tert-butyl hydroperoxide (t(26).
BHP) in studies with rats. These pigments decreased the
serum levels of alanine and aspartate aminotransferase and
SOURCES OF ANTHOCYANINS
reduced oxidative liver damage (20). In rats in which hepatic
Pigments of plant materials have called the attention of
scientists and the food industry in the last decades as source of
© Phcog.Net 2009 | www.phcog.net
129
Diego and Angela, Phcog Rev. 3(1):127-132 [2009]
extracts with biological properties. The coloration blue, red
and purple found in many fruits, vegetables and leaves are of
great interest since they are potential sources of anthocyanins
extracts. This interest has increased lately, since, these
pigments can be used as an alternative to artificial food
colorants and also, because they are bioactive compounds.
Anthocyanins functions in plants are similar to the general
functions of all flavonoids: antioxidant functions,
photoprotectors, defence mechanisms, and other ecological
functions (symbiotic phenomena). Since they give colour to
different parts of plants, they also play an interesting role in
the reproductive mechanisms: found in flowers, they serve to
attract pollinators and in seeds and fruits to attract seed
disseminators (27).
Anthocyanins pigments are usually extracted from plant
materials with an organic solvent. The most common is
ethanol and methanol containing a small amount of acid with
the objective of obtaining the flavylium cation form, which is
red and stable in a highly acid medium. However, acid may
cause partial hydrolysis of the acyl moieties in acylated
anthocyanins (28).
Recently, new techniques have been introduced for
anthocyanins extraction from different sources, such as
pressurized liquid extraction (29), sub and supercritical fluid
extraction (30), ultrasound assisted extraction, high hydrostatic
pressure or pulsed electric fields (31) and others.
The main sources of anthocyanins are red fruits, mostly
berries and red grapes, cereals, mainly the purple maize, and
vegetables (28, 32). Other potential sources of this
nutraceutical (Table 1) should be analyzed for commercial
proposes, since some plants can be found in great quantities
and the extraction of the bioactive compound is facilitated as
it is located in the external tissues of the plants.
Table 1 presents the concentration of anthocyanins from
various sources using different extraction methods employing
several solvents, and quantified by different anthocyanins
quantification methods. The wide range of anthocyanins
concentration obtained from the same source can be
associated to the different extraction methods and also to
different external and internal factors of the plant growth,
such as genetic and agronomic factors, intensity and type of
light, temperature and processing and storage of these
agricultural matters (17).
Environmental conditions are known to induce the
accumulation of anthocyanin pigments in the major groups of
higher plants, light and temperature factors being the most
studied ones. In the fruit of many crops such as grape, peach,
strawberry, eggplants and lychee, anthocyanin synthesis is
enhanced by sunlight and by cold weather (64, 65).
There are many other sources of anthocyanins-rich plants
around the world such as fruits (in general their skins),
flowers, stems, leaves and roots known and unknown until
now. It is interesting to note the relevance of some under
industrial utilized tropical fruits as potential source of
anthocyanins. One of these is the Brazilian fruit called
Jabuticaba (Myrciaria cauliflora).
© Phcog.Net 2009 | www.phcog.net
JABUTICABA AS A POTENTIAL SOURCE IN
BRAZIL
Jabuticabeira (Myrciaria cauliflora [Myrtaceae]) is a tree that
grows mainly in Brazil, most frequently in the states of São
Paulo, Minas Gerais, Rio de Janeiro, and Espírito Santo (66).
This specie is often cultivated in home gardens, small-scale
agricultural plots, or wild-harvested. They are primarily eaten
fresh and can be found in local markets; they are also used to
make jams, desserts, wines, liquors, and vinegars due to their
short shelf life, usually 3 to 4 days after harvesting the fruits
begin to ferment (7).
Jabuticaba is grape-like in appearance and texture, although its
skin is thicker and tougher. This fruit has a dark purple to
almost black skin color due to a high content of anthocyanins
(310-315 mg/ 100 g of fresh weight) that covers a white
gelatinous flesh inside (53, 67). It is 3 to 4 cm in diameter and
carries from one to four large seeds. The fruits are born
directly on the main trunks and branches of the plant, lending
to a distinctive appearance to the fruiting tree (68).
Even with few studies reported in the literature and a yet not
well known phytochemistry of this fruit, its sun-dried skins is
traditionally used as a treatment for hemoptysis, asthma,
diarrhea and chronic inflammation of the tonsils (69).
In a careful literature survey it was found only 7 articles that
linked the jabuticaba fruit to its anthocyanins pigments
published in journals indexed by the Web of Science and
Scopus database (Table 2). Of these, only 4 mentioned and/or
showed the potential of this Brazilian fruit as a source of
functional pigments (Table 2).
The first one (reference number 67 of Table 2) was published
in 1972 and its aim was to determine the concentration and
type of anthocyanins in the jabuticaba jam process. The
anthocyaninc extracts were purified and separated using thin
layer chromatography. The chromatography and chemical
properties (easy degradation) of these pigments supplied the
first indication about the structure of the two isolated
pigments (Peonidin and Peonidin-3-monoglucoside).
The other 2 papers (references 2 and 4 in Table 2) focused on
the quantitative analysis of anthocyanins from jabuticaba.
In reference number 68 more attention was given to the
identification of the anthocyanins present in jabuticaba and
their antiradical activities. A new depside, jaboticabin, together
with 17 known compounds were isolated from the jabuticaba
skin in this study.
The last 3 papers (references 70, 1, and 71 in Table 2) focused
on different aspects on the development of jabuticaba. The
first reference (70) discusses the accentuated increase of
anthocyanins concentrations at the end of the fruit growth
cycle. The second one (1) evaluates the antioxidant activity of
aqueous extract from jabuticaba. And the last paper (71) uses
the jabuticaba skin extracts as semiconductor sensitizer for
solar energy production. In this paper the results showed a
successful conversion of visible light into electricity by using
anthocyanins dye as wide band-gap semiconductor sensitizer
in dye-sensitized solar cells.
130
Diego and Angela, Phcog Rev. 3(1):127-132 [2009]
CONCLUSION
This review has summarized some important papers that
confirm that besides color, anthocyanins have properties that
are beneficial to human health, with potential physiological
effects such as anticancer, vasoprotective, anti-inflammatory,
hepatoprotective, among others.
Tropical under utilized fruits as jabuticaba (Myrciaria cauliflora)
in Brazil has demonstrated to be a good option of nonconventional sources of anthocyanins as natural food
colorings due to their dark purple skins rich in anthocyanins
(310-315 mg/ 100g of weight fresh). However, there is still a
lack of information in the literature (just 4 papers published in
journals indexed in the Web of Science and Scopus data) to
promote this Brazilian fruit as a potential source of functional
pigments.
ACKNOWLEDGEMENTS
The authors gratefully acknowledge the financial support of
CNPq (Conselho Nacional de Desenvolvimento Científico e
Tecnológico), Brazil.
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132
APÊNDICE II - DETALHAMENTO DO PROCEDIMENTO DE CÁLCULO DA
CONCENTRAÇÃO DE ANTOCIANINAS E FENÓIS
Cálculo da concentração de antocianinas
A concentração dos extratos antociânicos foi determinada pelo método do pH
diferencial, que é fundamentado nas transformações estruturais das antocianinas em função
do pH gerando soluções coloridas. O cátion flavílico, de coloração vermelha, é a forma
predominante em pH 1,0 enquanto que o carbinol, incolor, predomina em pH 4,5 (Figura
A1). Por isso, seguindo-se este método são feitas medidas espectrofotométricas de
antocianinas em soluções de pH 1,0 e 4,5, no comprimento de onda em torno de 500-550
nm (máximo de absorção das antocianinas) e 700 nm, para corrigir eventuais erros
referentes ao espalhamento da luz, já que os extratos podem apresentar suspensões
coloidais.
Figura A1 - Esquema das transformações estruturais das antocianinas em função do pH
gerando soluções coloridas.
201
Para se determinar o valor exato do comprimento de onda que resulta na
máxima absorção das antocianinas, primeiramente se é recomendado estudar o espectro de
varredura UV-vis (800 nm a 190 nm) para os extratos a serem analisados. Estes valores
conforme mencionado variam geralmente entre 465-550 nm, dependendo da fonte vegetal
utilizada. Para antocianinas de casca de jabuticaba foi-se encontrado 517 nm, já para
antocininas de jambolão 512 nm, matéria-prima que o grupo da professora Maria Angela de
Almeida Meireles também estuda como potencial fonte alternativa de antocianinas.
Foi observado que os espectros dos extratos ricos em antocianinas,
independente do solvente e do método de extração utilizado, apresenta pelo menos duas
bandas de absorção máxima de luz para os extratos, as quais se encontram em
aproximadamente 270-280 e 465-550 nm. Em alguns trabalhos é mencionado a presenca de
uma terceira banda ao redor de 200 nm, porém no presente trabalho somente foram
observados duas, uma em 275 e outra em 512 nm (Figura A2). Segundo a literatura o valor
de absorbância máxima entre 465 e 550 nm é exclusivamente dependente da concentração
de antocianinas, já o valor de absorbância máxima em torno de 275 nm é dependente
também de outros fatores, tais como outros compostos fenólicos: ácidos fenólicos, taninos,
dentre outros.
202
1.1
1
0.9
0.8
0.7
0.6
0.5
0.4
0.3
0.2
0.1
0
Soxhlet
Ultra-som
Abs
Percolação
200
250
300
350
400
450
nm
500
550
600
650
Figura A2 - Espectro de varredura UV-vis para os extratos de casca de jabuticaba obtidos
utilizando diferentes métodos de extrações.
Para o cálculo da concentração das antocianinas em extratos vegetais,
inicialmente, determina-se a absorbância “resultante” das leituras das soluções pela
Equação 1. Posteriormente, com a Equação 2 o cálculo da concentração das Antocianinas
Monoméricas Totais (AMT) presentes nos extratos é feito. A quantificação do teor de
antocianinas presentes em extratos contendo outra antocianina, que não a cianidina-3glicosídeo, como majoritária também pode ser feita pela Equação 2, embora os valores de
absortividade molar e massa molar sejam referentes a cianidina 3-glicosídeo (Ci-3-glu).
Embora, cianidina-3-glicosídeo seja utilizada como uma espécie de padrão de antocianinas,
devido à abundância desta antocianina na natureza, se o pesquisador conhece qual
antocianina é a majoritária em sua amostra o melhor é se calcular a concentração de
antocianinas em termos desta antocianina e não em termos de Ci-3-gli. Para se fazer este
cálculo é só substituir os valores de absortividade molar e massa molar na Equação 2.
203
A = (A max −A 700 ) pH1, 0 − (A max − A 700 ) p 4,5
AMT (mg / L) = ( A x PM x FD x1000) / (ε x l )
(1)
(2)
Sendo Amax - absorbância no máximo comprimento de onda, A700 - absorbância
a 700 nm, PM - Peso Molecular, FD - Fator de Diluição, ε - absortividade molar (para
cianidina-3-glicosídeo ε = 26,900 L/mol.cm) e l - comprimento do percurso ótico da cubeta
(para a cubeta de vidro utilizada neste trabalho l = 1 cm).
Além de quantificar a concentração de antocianinas em extratos vegetais o
método descrito pode ser aplicado na determinação do teor destes compostos em amostras
como bebidas, doces, etc.
Neste trabalho etanol e água acidificada foram utilizados como solventes nos
processos de extração de antocianinas de cascas de jabuticaba. Como a evaporação do
solvente etanol é um procedimento fácil (através de rotaevaporador) optou-se em todos os
experimentos envolvendo etanol proceder com a eliminação do solvente para obtermos um
extrato bruto o qual seria, então diluído em água e caracterizado em termos de antocianinas
através do método espectofotométrico descrito acima. Devido à dificuldade de eliminação
do solvente água através dos utensílios disponibilizados no laboratório a solução aquosa
contendo antocianinas extraídas foi diluída também em água e caracterizada em termos de
antocianinas diretamente.
Uma vez que um dos objetivos deste estudo foi comparar diferentes métodos de
extração para a obtenção de extratos ricos em antocianinas padronizou-se expressar os
resultados em termo de mg de cianidina-3-glicosídeo/g de casca de jabuticaba seca.
Utilizando-se água o procedimento de cálculo foi mais simples, uma vez que somente é
necessário multiplicar o valor encontrado de AMT (mg Cianidina-3-glicosíceo/L) pelo
volume de água em litros utilizado e dividir pela massa de matéria-prima seca em gramas
(Equação 3). Já utilizando etanol o procedimento de cálculo teve duas etapas: i) Dividir o
valor encontrado de AMT (mg Cianidina-3-glicosíceo/L) pela concentração do extrato
bruto diluído em água inicialmente em mg de extrato/L (em todos os experimentos
204
adotamos diluir 20 mg de extrato bruto (após evaporação do etanol) em 10 mL de água –
0.002 mg de extrato/L) (Equação 4). Posteriormente multiplicou-se este valor pelo massa
de extrato bruto obtida em miligramas e dividiu-se pela massa de matéria-prima seca em
gramas.
mg Ci − 3 − gli / g de material sec o =
AMT (mg Ci − 3 − gli / L) x V ( L)
(3)
m( g de cascas de jabuticaba sec a)
mg Ci − 3 − gli / mg de extrato =
mg Ci − 3 − gli / g de material sec o =
AMT (mg Ci − 3 − gli / L)
(4)
(mg de extrato / L)
AMT ( mg Ci − 3 − gli / L ) x mg de extrato
(5)
( mg de extrato / L ) x g de cascas de jabuticaba sec a
Cálculo da concentração de compostos fenólicos
A concentração de compostos fenólicos totais nos extratos foi estimada
utilizando o método de Folin–Ciocalteau, método este baseado na reação colorimétrica de
oxidação/redução do ácido fosfomolíbdico-fosfotúngstico pelas hidroxilas fenólicas,
originando óxidos azuis de tungstênio (W8O23) e de molibidênio (Mo8O23). Um complexo
de coloração azul-esverdeada que absorve luz em comprimentos de onda entre 620 e 770
nm, com um comprimento de onda máximo ao redor de 725 nm. A reação ocorre em meio
alcalino e a solução de carbonato de sódio é a mais utilizada.
Neste trabalho o teor de fenóis totais foi calculado com base na curva de
calibração (Figura A3), previamente construída, de ácido gálico (AG) e foram expressos
como mg de ácido gálico equivalentes (AGEs)/g de material seco. O procedimento de
cálculo para obtermos a concentração de compostos fenólicos como mg de ácido gálico
equivalentes (AGEs)/g de material seco foi bem similar ao adotado para se determinar o
conteúdo de antocianinas nos extratos previamente descrito. A Equação 6 foi utilizada para
se determinar a concentração de compostos fenólicos presentes nos extratos de casca de
jabuticaba utilizando água como solvente, onde FD é o fator de diluição. O valor de FD
205
oscilou entre 5 a 20, dependendo da concentração de compostos fenólicos para o valor de
absorbância ficar ente 0-1 (dentro da faixa confiável da curva de calibração (obedecendo a
lei de Beer), previamente construída, de ácido gálico (mg de AG/mL). Equações 7 e 8
foram utilizadas quando etanol foi o solvente escolhido. Em todos os experimentos
utilizando etanol como solvente adotou-se diluir 10 mg de extrato bruto (após evaporação
do etanol) em 10 mL de água – 1 mg de extrato/mL.
y = 0,08035x + 0.0105
R2 = 0.997
0.12
mg de AG/mL
0.1
0.08
0.06
0.04
0.02
0
0
0.2
0.4
0.6
0.8
1
1.2
Absorbância
Figura A3 - Curva de calibração previamente construída de ácido gálico (AG)
mg AGEs / g de material sec o =
(mg AGEs / mL) x FD x V(L)
m(g de cascas de jabuticaba sec a )
mg AGEs / mg de extrato =
mg AGEs / g de material sec o =
(mg AGEs / mL)
(mg de extrato / mL)
(6)
(7)
(mg AGEs / mL) x mg de extrato
(mg de extrato / mL) x g de cascas de jabuticaba sec a
206
(8)
APÊNDICE III - ARTIGO DE REVISÃO - CAROTENOID PIGMENTS
ENCAPSULATION: FUNDAMENTALS, TECHNIQUES AND RECENT TRENDS
Publicado no periódico The Open Chemical Engineering Journal.
207
208
42
The Open Chemical Engineering Journal, 2010, 4, 42-50
Open Access
Carotenoid Pigments Encapsulation: Fundamentals, Techniques and
Recent Trends
Diego T. Santos and M. Angela A. Meireles*
LAboratory of Supercritical Technology: Extraction, Fractionation, and Identification of vegetable extracts (LASEFI),
Department of Food Engineering (DEA), University of Campinas (UNICAMP), Rua Monteiro Lobato, 80, Cid.
Universitária Zeferino Vaz, 13083-862, Campinas - SP, Brazil
Abstract: Supercritical fluids have become an attractive alternative due to environmentally friendly solvents. The methods that use supercritical fluids can be conveniently used for various applications such as extraction, reactions, particle
formation and encapsulation. For encapsulation purposes, the processing conditions given by supercritical technology
have important advantages over other methods that include harsh treatments with regard to pH, temperature, light, the use
of organic solvents, etc. Unstable functional pigments such as carotenoids extracted from natural sources have been encapsulated to overcome instability problem. Thus, the most used techniques applicable to this intention are described and
discussed in this review as well the recent advances and recent trends in this topic that involves the use of supercritical
fluids.
Keywords: Encapsulation, unstable functional pigments, carotenoids, supercritical fluids.
INTRODUCTION
Encapsulation is defined as a technology of packaging
solids, liquids, or gaseous materials in matrices that can release their contents at controlled rates under specific conditions. The encapsulation technology has been used by the
food industry for more than 60 years [1].
Protective encapsulation of food ingredients, enzymes,
cells or other functional compounds may be thus achieved in
small capsules of different nature. The main aim of encapsulation in the food area is to protect sensitive food components from moisture, oxidation, heat, light or extreme conditions during processing in an effort to increase their shelf
life, or to mask component attributes, as undesirable flavours, to meet consumer requests for organoleptic quality
and functionality. Furthermore, an optimized design of capsules, choosing the best encapsulation process and conditions, might provide a controlled release of active compounds during processing or storage [2].
In the last years, encapsulation has received increasingly
growing attention resulting in a great number of applications
in industry, agriculture, medicine, pharmacy and biotechnology. Diverse techniques have been studied and employed to
form the desired capsules, including spray drying, liposome
entrapment, coacervation, gelation, emulsion phase separation, etc [3]. Some of the most used techniques are described
and discussed in this review. In addition, this paper gives an
overview of the recent advances in the various encapsulation
techniques applicable for stabilizing unstable functional
pigments, such as, carotenoids from natural sources. Finally
*Address correspondence to this author at the laboratory of Supercritical
Technology: Extraction, Fractionation, and Identification of vegetable extracts (LASEFI), Department of Food Engineering (DEA), University of
Campinas (UNICAMP), Rua Monteiro Lobato, 80, Cid. Universitária Zeferino Vaz, 13083-862, Campinas - SP, Brazil; Tel: 00551935214033;
E-amil: [email protected]
1874-1231/10
a discussion about the recent trends in this topic that involves
the use of supercritical fluids instead of conventional solvents will be debated.
ENCAPSULATION CONCEPT
In general the purpose of encapsulation is to protect its
contents from the environment which can be destructive
while allowing small molecules to pass in and out of the
membrane. According to Jizomoto and coworkers the encapsulation concept has its origin in the idealization of the cell
model, in which the nucleus is involved by a semi-permeable
membrane that protects it from the external medium and also
controls the entrance and exit of substances. Other natural
examples include birds and reptile egg shells, plant seeds,
fruit and vegetable skins, seashells, etc [4].
The encapsulation process involves coating or entrapment of a material, usually a liquid but can be a solid or gas,
or a mixture into another material. This material is also
known as the core material. The coating material can also be
called the capsule, wall material, membrane, carrier or shell
[5].
The first applications of encapsulation were done in 1954
in the photography industry to produce pressure-sensitive
dye capsules for the manufacturing of carbonless copying
paper, since then, along the last decades encapsulation processes were developed and used in a variety of industries [6,
7]. In the food industry, diverse encapsulation techniques
have been applied to protect unstable compounds, such as,
flavours, pigments, vitamins, enzymes, microorganisms, and
others [8].
ENCAPSULATION
PIGMENTS
OF
UNSTABLE
FUNCIONAL
Epidemiological evidences suggest that a diet rich in
fruits and vegetables plays an important role in reducing the
2010 Bentham Open
Carotenoid Pigments Encapsulation
Table 1.
The Open Chemical Engineering Journal, 2010, Volume 4
43
Encapsulation Processes Division Forms Adopted by Some Reseachers
Number of
Categories
Criterion
3
According to the dominant structural component
used in their assembly
Categories
References
1) Surfactant-based (emulsion polymerization)
2
According to the encapsulation process
3
According to the encapsulation technique. All
encapsulation techniques are modifications of three
basic techniques
2) Lipid-based (emulsion phase separation)
3) Biopolymer-based
(inclusion
coacervation, gelation, spray coating)
[14]
complexation,
1) Chemical processes (coacervation, cocrystallization,
inclusion complexation, emulsion polymerization)
2) Mechanical processes (spray coating, fluidized bed
coating)
[6]
1) Phase separation (coacervation)
2) Spray drying
[18]
3) Solvent extraction/evaporation
1) Physical nature (spray coating, extrusion coating,
centrifugal and rotational suspension separation,
fluidized bed coating, liophilization, cocrystallization)
3
According to the nature of the combination between
coating or entrapment material and core material
2) Chemical nature (inclusion complexation, emulsion
polymerization)
[8]
3) Physical-chemical nature (coacervation, emulsion
phase separation, liposome entrapment)
incidence of many diseases. The reason of this accomplishment can be the coloration of these foods, in general, a rich
source of many antioxidants [9].
The increasing concern of consumers over the use of synthetic additives in products has pushed the food, pharmaceutical and cosmetic industries toward replacing these synthetic
additives by natural products. However, difficulties may be
encountered due to the instability of these compounds. An
example are natural pigments with functional properties such
as carotenoids, the most common group of pigments in nature [10].
In order to overcome the instability problem of these bioactive compounds, which results in restricted commercial
applications, encapsulation has become an important tool,
helping to increase shelf life and protecting the biological
properties of the material [11]. Moreover encapsulation allows the controlled release of these functional pigments under desired conditions, for instance when ingested in the
body [5].
CAROTENOIDS
Carotenoids significance is not only due to their colorant
property, they also are very important for health. This compound is source of vitamin A and a precursor of important
chemicals responsible for the flavor of foods and the fragrance of flowers. It shows important biologic activities associated with antioxidant properties, such as strengthening
the immune system, decreasing the risk of degenerative illnesses such as cancer, preventing the risk of cardiovascular
disease, preventing macular degeneration, and reducing the
risk of cataracts [12, 13].
Carotenoid pigments are a diverse group of lipophilic
compounds that contribute to the yellow to red colors of
many foods. They are polyenes consisting of 3 to 13 conju-
gated double bonds and in some cases 6 carbon ring structures at one or both ends of the molecule. The most common
carotenoid types are -Carotene, lycopene, lutein, and zeaxanthin [14].
Unstable carotenoids have their oxidative degradation
triggered by light, temperature and/or extreme pH in the
presence of oxygen. To improve their stability to oxidation
different encapsulation techniques have been studied, resulting in a relative success and increasing additionally the dissolution rate of these lipossoluble compounds in water [15,
16].
ENCAPSULATION TECHNIQUES
Diverse techniques have been studied and employed to
form the capsules in different industries, including spray
coating: spray drying, spray chilling, spray cooling; extrusion coating; fluidized bed coating; liposome entrapment;
simple and complex coacervation; inclusion complexation;
emulsion polymerization; centrifugal and rotational suspension separation; thermal and ionic gelation; emulsion phase
separation; liophilization; cocrystallization; etc [1, 3, 17].
For convenience some scientists have divided the structured delivery systems and, consequently, the encapsulation
processes in different forms. Table 1 shows the most frequently division forms adopted.
In this review paper will be assumed that the capsules
can be prepared by various techniques, which feature that
partly competing, partly complementary characteristics and
many encapsulation processes are modifications of basic
conventional techniques. Due to its high coverage the division form based on the nature of the combination between
coating or entrapment material (called also as wall material
or carrier material) and core material will be adopted in this
review. Then, the three encapsulatipon categories: based on
44 The Open Chemical Engineering Journal, 2010, Volume 4
Physical nature, Chemical nature and Physical-chemical nature will be described and discussed below in order to demonstrate the recent advances/modification in some of the
various encapsulation techniques that can be applicable for
stabilizing unstable carotenoid pigments.
ENCAPSULATION
TECHNIQUES
BASED
ON
PHYSICAL NATURE COMBINATION BETWEEN
COATING MATERIAL AND CORE MATERIAL
Among the encapsulation physical methods, some examples are: spray coating: spray drying, spray cooling/chilling;
extrusion coating, centrifugal and rotational suspension separation, fluidized bed coating, liophilization, cocrystallization,
etc [8].
In this paper, due to their higher aplicability especial attention will be given to the encapsulation techniques spray
coating and spray drying. In the section Advantages and
Limitations of the Conventional Encapsulation Methods Applicable for Stabilizing Carotenoid Pigments the other techniques will be rapidly discussed.
Spray Coating
The spray coating process involves the dispersion of the
substance to be encapsulated in a coating or entrapment material, followed by atomization and spraying of the mixture
into a chamber. When the atomization is done when a hot
fluid desiccant (air) is passing into the chamber the spray
coating process is called by spray drying and when the air
desiccant is cold the process is called by spray cooling/chilling. After pass to the chamber the resulting capsules
are then transported to a cyclone separator for recovery [19].
Another difference between spray drying and spray cooling/chilling is that in the last one there is generally no water
to be evaporated. There is emulsification of the compounds
into molten wall materials, followed by atomization to disperse droplets from the feedstock [20]. Although spray cooling and spray chilling are not the same technique, differing
slightly only in the vessel temperature in which the coating
material is sprayed [21], due to their similarity in this work
they were grouped.
Spray drying is a commercial process widely used in
large-scale production of encapsulated compounds. The
merit of the process is due to several factors: high availability of equipment, low process cost, wide choice of coating or
entrapment material, good encapsulation efficiency, good
stability of the finished product, and large-scale production
in continuous mode [22].
Spray drying is relatively simple and of high throughput
but should not be used for highly temperature-sensitive compounds. To this kind of compounds, such as vitamins, enzimes, flavours, pigments, essential oils and others, spray
cooling/chilling may be an appropriate method [8, 19].
Moreover, control of the particle size in the spray coating is
difficult, and yields for small batches are moderate [23].
Extrusion Coating
According to Madene and coworkers the two major encapsulation industrial processes are spray drying and extrusion coating [6]. However, to food compounds the encapsu-
Santos and Meireles
lation by extrusion is a relatively new process compared to
spray drying [1].
The advantage of this method is that the material is totally surrounded by the wall material and that any residual
core is washed from the outside. Owing to this benefit this
encapsulation method is called by some authors true or glass
encapsulation [3].
Extrusion coating can be organized in two types: simple
extrusion and centrifugal extrusion. In general, the process
involves forcing a core material in a molten wall material
mass through a die (laboratory scale) or a series of dies of a
desired cross-section into a bath of desiccant liquid. The
coating material hardens on contacting the liquid, entrapping
the active substances. Then the extruded filaments are separated from the liquid bath, dried, and sized [12].
Basically, the difference between simple extrusion and
centrifugal extrusion procedures is that in the last one the
nozzle used is not a simple nozzle (the nozzle has a coaxial
opening). Centrifugal extrusion is a liquid coextrusion process utilizing nozzles consisting of concentric orifice located
on the outer circumference of a rotating cylinder. The core
and the carrier materials are fed through, respectively, the
inner and outer opening. The core and the coat fluids must be
immiscible. At the tip of the coaxial nozzle the two fluids
form a unified jet flow by centrifugal force, which is responsible to form the droplets [24].
In order to optimize the centrifugal extrusion coating
some researchers are studying the recycle of the excess coating fluid from the centrifugal extrusion, while the resulting
capsules are hardened. This modified process has been called
by recycling centrifugal extraction [25]. However, the major
problem of this method is related to the difficulty of obtaining capsules in extremely viscous carrier material melts [26].
ENCAPSULATION
TECHNIQUES
BASED
ON
CHEMICAL NATURE COMBINATION BETWEEN
COATING MATERIAL AND CORE MATERIAL
According Jackson and Lee (1991) there are only two encapsulation chemical methods: inclusion complexation and
emulsion polymerization [8]. The last one, also called by
interfacial polymerization, will not be presented in this paper, spite of it been a well established method it is most used
to trap inorganic particles and not necessarily improve their
stability [27].
Inclusion Complexation
Inclusion complexation or molecular inclusion is called
the encapsulation process that uses -cyclodextrin molecular
conformation to entrap any core material. A typical application is the protection of unstable and high added value specially flavours [28], but recently diverse bioactive compounds have been encapsulated by this technique [29].
The external part of the -cyclodextrin molecule is hydrophilic, whereas the internal part (central cavity) is hydrophobic. Some particular molecules, which are apolar and
have suitable molecular dimensions to fit inside the cavity,
can be entrapped through a hydrophobic interaction. The
mechanism involved in this method is based on the replacement of water molecules by less polar molecules [30].
Carotenoid Pigments Encapsulation
Comparing the inclusion complexation procedure to form
capsules to the others mentioned before similarities can be
observed. The core material is diluted in the coating material
(-cyclodextrin) also, but the -cyclodextrin needs to be dissolved first in water to form an aqueous solution to then be
mixed to the core material. Afterwards, an inclusion complex
is formed between -cyclodextrin and core material, being
the precipitate recovered and dried by conventional means
[20].
To eliminate the last required step: capsules drying less
amount of water can be utilized to disperse the coating material avoiding additional steps and consequently reducing
costs [31].
In spite of important progress to improve economical aspects of the encapsulation by inclusion complexation, such
as reducing the water content in the final product, still the
relatively expensive price of -cyclodextrins and the undesirable release of the formed complex into the mouth are
barriers to the industrial development of this technique [3,
21].
ENCAPSULATION
TECHNIQUES
BASED
ON
PHYSICAL-CHEMICAL NATURE COMBINATION
BETWEEN COATING MATERIAL AND CORE MATERIAL
Some examples of the encapsulation physical-chemical
methods are: coacervation, emulsion phase separation,
liposome entrapment, etc [8]. Coacervation technique is
widely used in the industry, therefore it will be described
below while the others techniques will be briefly discussed
in the section Advantages and Limitations of the Conventional Encapsulation Methods.
Coacervation
Coacervation is often regarded as the original method of
encapsulation [20]. This technique was the first encapsulation process studied and was initially employed by Green
and Scheicher in 1955 to produce capsules for the manufacturing of carbonless copying paper as mentionated before.
Coacervation technique is the separation into two liquid
phases in colloidal systems. The phase more concentrated in
colloid component is the coacervate (a polymer rich phase)
and the other phase is the equilibrium solution (almost
polymer free) [32]. Coacervation in aqueous systems is subdivided into simple and complex coacervation.
In the first procedure step, a three-phase system consisting of a liquid manufacturing vehicle phase (solvent), a core
material phase, and a coating material phase (formed by one
polymer – simple coacervation; by two polymers – complex
coacervation) is formed by a direct addition. Afterwards the
deposition of the liquid polymer coating around the core
material by electrostatic attraction is initialized by controlled
physical mixing of the coating material and the core material
in the manufacturing vehicle in the liquid phase. After accomplished the deposition it is necessary the stabilization
employing toxic chemical agents, such as glutaraldehyde [1].
Due to the ineffective last step required, according Thomasin and coworkers, coacervation is frequently impaired by
residual solvents and coacervating agents found in the
spheres. Furthermore, it is not well suited for producing
The Open Chemical Engineering Journal, 2010, Volume 4
45
spheres in the low size range. Additionally, others drawbacks
about this method that is necessary to mention is its very
expensive and rather complex process [33].
ADVANTAGES AND LIMITATIONS OF THE CONVENTIONAL ENCAPSULATION METHODS APPLICABLE FOR STABILIZING CAROTENOID PIGMENTS
Table 2 summarizes various advantages and limitations
of some conventional encapsulation methods applicable for
stabilizing unstable carotenoids.
Recently alternative technologies and/or even some small
modification as change the conventional organic solvent to
“greener solvent” such as supercritical fluids have been
evaluated to eliminate some limitations of the conventional
encapsulation methods [34].
Advantages as elimination of organic solvents, encapsulation under mild conditions and formation of homogeneus
micro-nanocapsules can be achieved using supercritical fluid
based-techniques. These techniques could avoid carotenoid
fast degradation [35]. Thus, the recent trends in this topic
will be discussed.
ENCAPSULATION USING SUPERCRITICAL FLUIDS
In the recent years, new encapsulation techniques utilizing supercritical fluids have been developed in order to overcome some of the disadvantages of the conventional techniques [49]. Some of these drawbacks are: a) poor control of
particle size and morphology; b) degradation of thermo sensitive compounds; c) low encapsulation efficiency; d) low
yield [50].
The use of supercritical fluids as phase separating agents
has been intensively studied also to minimize the amount of
potentially harmful residues in the capsules and most effectively control [51, 52].
According to Cocero and coworkers carbon dioxide
(CO2) is most solvent used for encapsulation purposes due to
the supercritical region can be achieved at moderate pressures and temperatures (Tc = 304.2 K, Pc = 7.38 MPa);
therefore, working with supercritical CO2 it is possible to
carry out the process at near-ambient temperatures, avoiding
the degradation of thermolabile substances [53].
Several encapsulation processes that use supercritical fluids have been developed. These processes can be classified
according to the role of the supercritical fluid in the process:
solvent [Rapid Expansion of Supercritical Solutions
(RESS)]; Supercritical Solvent Impregnation (SSI), solute
[Particles from Gas Saturated Solutions (PGSS)] or antisolvent [Supercritical Anti-Solvent (SAS); Supercritical
Fluid Extraction of Emulsions (SFEE)] [54].
Given that the two most commonly encapsulation methods employed using supercritical fluids are the Rapid Expansion of Supercritical Solution (RESS) and the Supercritical
Anti-Solvent (SAS) methods [54], particular consideration
will be given to these techniques. Furthermore the potential
application of supercritical fluids in the micro-nanoencapsulation technology with emulsions called by the literature of
Supercritical Fluid Extraction of Emulsions (SFEE) will be
presented.
46 The Open Chemical Engineering Journal, 2010, Volume 4
Santos and Meireles
Table 2. Advantages and Limitations of Some Conventional Encapsulation Methods Applicable for Stabilizing Unstable Carotenoids
Conventional
Encapsulation
Method
Principle
Advantages
Limitations
References
Dispersion of the core material in a entrapment material, followed by atomization
and spraying of the mixture in a hot air
desiccant into a chamber
Low process cost; wide
choice of coating material;
good encapsulation efficiency; good stability of the
finished product; possibility
of large-scale production in
continuous mode
Can degradate highly temperature-sensitive compounds;
control of the particle size is
difficult; yields for small
batches are moderate
[19, 20, 22,
23]
The same of the spray drying differing only
that the air desiccant is cold
Temperature-sensitive compounds can be encapsulated
Difficult control of the particle
size; moderate yields for small
batches; special handling and
storage conditions can be required
[21, 23, 36]
Simple Extrusion
Forcing a core material in a molten wall
material mass through a die (laboratory
scale) or a series of dies of a desired crosssection into a bath of desiccant liquid. The
coating material hardens on contacting
liquids, entraping the active substances
The material is totally surrounded by the wall material;
any residual core is washed
from the outside; it is a relatively low-temperature entrapping method
The capsule must be separated
from the liquid bath and dried;
is difficult to obtain capsules in
extremely viscous carrier material melts
[3, 12, 22, 26]
Centrifugal Extrusion
Similar of simple extrusion differing that
the core material and coating material form
a unified jet flow only at the end through a
nozzle with a coaxial opening (coextrusion) by centrifugal force
The same of simple extrusion
The same of simple extrusion
[24]
Ionic Gelation
Coating material with dissolved core material is extruded as drops within an ionic
solution. The capsules are formed by ionic
interaction
Organic solvents and extreme
condions of temperature and
pH are avoided
Mainly used on a laboratory
scale; the capsules, in general,
have high porosity which promotes intensive burst
[37, 38]
Thermal Gelation
The principle is almost the same of ionic
gelation’ principle, nonetheless there is no
necessity of an ionic solution to form a
gelled drop, the gelation is only due to
thermal parameters
The same of ionic gelation
The same of ionic gelation
[37, 38]
Fluidized Bed Coating
This technique relies upon by nozzle spraying the coating material into a fluidized bed
of core material in a hot environment
Low cost process; it allows
specific capsule size distribution and low porosities into
the product
Degradation of highly temperature-sensitive compounds
[28, 39]
Lyophilization/Freeze
Drying
The entrapment occurs by lyophilization of
an emulsion solution contaning a core
material and a coating material
Thermosensitive substances
that are unstable in aqueous
solutions may be efficiently
encapsulated by this technique
Long processing time; expensive process costs; expensive
storage and transport of the
capsules
[40]
Inclusion Complexation
Particular apolar molecules are entrapped
through a hydrophobic interaction inside
the -Cyclodextrin cavity replacing water
molecules
Very efficient to protect
unstable and high added
value apolar compounds such
as flavours
Encapsulation restricted to
apolar compounds with a suitable molecular dimensions; cyclodextrin price is expensive;
frequently undesirable release
of the formed complex
[3, 21, 28, 30]
Emulsion Polymerization
Core material is dissolved into polymerization sollution. The monomers are polymerized to form capsules in an aqueous solution
Micro-nanocapules with
narrow size distribution can
be obtained
Difficult control of the capsule
formation (polymerization)
[41, 42]
Spray Drying
Spray
Cooling/Chilling
Carotenoid Pigments Encapsulation
The Open Chemical Engineering Journal, 2010, Volume 4
47
Table 2. contd….
Conventional
Encapsulation
Method
Principle
Advantages
Limitations
References
[1, 33, 43, 44]
Coacervation
The entrapment is due to the deposition of
a liquid coating material around the core
material by electrostatic attraction
Can be used to encapsulate
heat-sensitive ingredients due
to done at room temperature
Toxic chemical agents are
used; the complex coacervates
are highly unstable; there are
residual solvents and coacervating agents on the capsules
surfaces; spheres low size
range; expensive and complex
method
Emulsion Phase Separation
The core material is added in the polar or
apolar layer of an oil-in-water emulsion O/W or water-in-oil - W/O emulsion. The
emulsions are prepared using a surfactant
Polar, non-polar (apolar), and
amphiphilic can be incorporated; emulsions can either be
used directly in their “wet”
state
Instable when exposed to environmental stresses, such as
heating, drying, etc; limited
number of emulsifiers that can
be used
[45, 46]
Liposome Entrapment
Phospholipids are dispersed in an aqueous
phase spontaneously formation a liposome.
A core material is entrapment into a
liposome
Either aqueous or lipidsoluble material can be encapsulated; suitable to high
water activity applications;
efficient controlled delivery
Mainly used on a laboratory
scale
[47, 48]
RAPID EXPANSION OF SUPERCRITICAL SOLUTION (RESS) METHOD
In the RESS method the solution of core material plus
coating material is solubilized in a supercritical fluid and the
solution is expanded rapidly through a nozzle. Thus, the solvent power of supercritical fluid dramatically decreases and
eventually occurs the co-precipitation of both substances
[55]. According to Mishima and Matsuyama (2006) it is difficult to disperse the core material homogenously in the coating material in the absence of surfactants or a high shear
condition in the supercritical CO2 [56]. In order to avoid this
homogeneity problem mechanical agitation or ultrasonic
irradiation apparatus have been added into the high-pressure
vessel [56, 57].
RESS technique is environmental friendly because the
capsule is completely solvent free. Unfortunately, most wall
materials exhibit little or no solubility in supercritical fluids,
limiting this technique to restrict applications [55].
To overcome the low solubility limitation of the wall materials in CO2, alternative organic supercritical solvents has
been employed. Another procedure by modification the
original RESS process has been carried out also for eliminating this solubility problem. This modified process has been
named by RESS-non-solvent process (RESS-N). In this
process, a liquid antisolvent for the coating material is used
as a cosolvent for improving the solubility in the supercritical fluid [58].
According to Cocero and coworkers besides the solubility limitations, another major problem of RESS techniques is
the difficulty to control the morphology and loading of the
capsules [53].
SUPERCRITICAL ANTI-SOLVENT (SAS) METHOD
The encapsulation by SAS technique, also called by Gas
Anti-Solvent (GAS) method, is based on the same simple
principle of RESS method whereby a core material and a
carrier are co-precipitated together. In SAS method the antisolvent (non-solvent) property of supercritical carbon dioxide (CO2) is used, since most wall materials and core material are not soluble in supercritical CO2 [49].
The basic principle of the SAS method is based on a
rapid decrease in the solubilization power of a solvent by
addition of a second solvent as antisolvent. Upon mixing, the
supercritical fluid (antisolvent) saturates the conventional
liquid solvent and depletes it by extraction. Particle size distribution can be partially controlled by adjusting the values
of temperature, pressure and composition. The high viscosity
of the coating material-CO2 solutions during atomization or
the solvent extraction process, generally, leads to inconsistency of the particle size, strong particle agglomeration, and
also incomplete encapsulation [59].
Another disadvantage of the supercritical antisolvent
process is the difficulty to remove the remaining solvent
completely because the process generally carries out in a
batch discontinuous process [59].
In order to overcome some of the disadvantages mentioned above Chattopadhyay an coworkers have developed
and patented a new encapsulation method called Supercritical Fluid Extraction of Emulsions (SFEE). Essentially this
method is combines the flexibility of particle engineering
using different emulsion systems with the efficiency of large
scale, continuous extraction ability, provided by supercritical
fluids [52].
According to Perrut and coworkers the application of supercritical fluids in the encapsulation technology with emulsions appears as a natural decision to avoid the main problems of each technology separately. Emulsion phase separation, emulsion polymerization and others emulsion techniques usually involve large quantities of organic solvents,
and the removal of them involves additional separation proc-
48 The Open Chemical Engineering Journal, 2010, Volume 4
Santos and Meireles
Table 3. Articles Published in Journals Indexed in the Web of Science and Scopus Databases About Carotenoid Encapsulation Using
Supercritical Fluids
Article Title
Encapsulation Method
References
1
Carotenoid processing with supercritical fluids
SAS and SEDS
[66]
2
Precipitation of -carotene and PHBV and co-precipitation from SEDS technique using
supercritical CO2
SEDS
[50]
3
Precipitation of lutein and co-precipitation of lutein and poly-lactic acid with the
supercritical anti-solvent process
SAS
[68]
4
Co-precipitation of -carotene and polyethylene glycol with compressed CO2 as an
antisolvent: effect of temperature and concentration
SAS
[69]
5
Production of natural carotene-dispersed polymer microparticles by SEDS-PA
co-precipitation
SEDS
[64]
6
Co-precipitation of carotenoids and bio-polymers with the supercritical anti-solvent process
SAS
[17]
7
Requirements for non-food applications of pea proteins. A Review
SFEE
[35]
ess and the use of high temperatures. On the other hand, supercritical fluids regularly are not able to produce capsules
below the micrometer range or the products obtained present
agglomeration problems [60].
According Cocero and coworkers the difference from
SAS processes and the SFEE, among others, is that an emulsion containing the core materials to be precipitated dissolved in its dispersed phase (conventional liquid solvent) is
injected instead of injecting a simple solution of the core
materials. Effectively during the “extraction” (encapsulation)
it can be expected that first the droplets of the disperse phase
become saturated by CO2, and then the solvent is extracted
by CO2 from them. Therefore during the saturation with CO2
each droplet behaves as a miniature SAS Anti-Solvent precipitator [53].
ENCAPSULATION OF CAROTENOID PIGMENTS
USING SUPERCRITICAL FLUIDS
Due to the presented potential advantages of the encapsulation process using supercritical fluids for carotenoid encapsulation these techniques seem to be more appropriate.
In a careful literature survey in Web of Science and Scopus databases it was found several articles about carotenoid
pigments encapsulation. But, more specifically about carotenoid encapsulation using supercritical fluids this amount
was reduced intensively to only 7 articles (Table 3).
Conventional encapsulation techniques such as spray
drying and inclusion complexation were the most common
methods evaluated for encapsulating carotenoids. Other studies, aiming indirectly at the development of alternative technologies towards encapsulating carotenoids in wall materials
using supercritical technology were found, but the amount
was still scarce. Even though, a significant increase in this
amount could be observed since 2006 until now (Fig. 1). Fig.
1 also shows that approximately 80% of the articles that aims
for developing supercritical fluids based technologies towards encapsulating carotenoids have been published in the
last 4 years.
Analyzing the literature about the evolution of encapsulating technologies carotenoids using supercritical fluids
Years 2002-2004
13%
Years 2004-2006
6%
Years 2008-2009
43%
Years 2006-2007
38%
Fig. (1). Distribution of the published articles that aims for
developing supercritical fluids based technologies towards
encapsulating carotenoids.
could be observed that it can be divided in four sequencial
parts. 1) First, a complex study of the precipitation of many
solutes, including carotenoids, from supercritical fluids by
rapid expansion (RESS process) was done in order to avoid
thermal decomposition that generally occurs by milling
process [61]; 2) After that, carotenoid precipitation from
liquid solvents, with dissolution by high-pressure or supercritical CO2 as an antisolvent to create supersaturation (GAS
or SAS processes, respectively) was evaluated as a crystallization process [62]; 3) In this part, the co-precipitation of
many solutes such as carotenoids and coating materials towards protect and stabilize them was extensively studied [63,
64]; 4) Finally, carotenoid encapsulation by RESS, GAS and
SAS techniques have been better studied (being in some
cases also successfully scaled up) and other techniques such
as the novel Solution Enhanced Dispersion by Supercritical
(SEDS) fluids, based on the principle of SAS, has been applied (Table 3) [64-67].
In SEDS process, a nozzle with two coaxial passages allows introducing supercritical CO2 and a solution with core
and coating materials into the particle formation vessel
where pressure and temperature are controlled. The high
velocity of supercritical CO2 allows breaking up the solution
into very small droplets since that the velocity is set up to
Carotenoid Pigments Encapsulation
extract the solvent from the solution (SAS principle) at the
same time as it meets and disperses the solution [65].
The use of a coaxial nozzle for encapsulation purposes is
not new. In this review paper was demonstrated that in extrusion coating method the same nozzle type was used to the
same purpose [24]. Thus, it can be seen that the recent encapsulation methods are based on modifying conventional
techniques.
CONCLUSION
This review paper has demonstrated the potentiality of
the use of supercritical fluids based encapsulation to protect
and stabilize unstable pigments. Among the advantages are:
form micro- or even nanoparticles with narrow particle distribution, reduce or even eliminate the residual organic solvent in the product, and control product quality.
Nowadays alternative technologies based on small modification as change the conventional organic solvent to
“greener solvent” such as supercritical fluids have been
evaluated successfully to eliminate some limitations of the
conventional encapsulation methods.
ACKNOWLEDGEMENTS
The authors are grateful to CNPq for the doctorate fellowship (141894/2009-1) and for the financial support
(580401/2008-1).
The Open Chemical Engineering Journal, 2010, Volume 4
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Revised: November 26, 2009
Accepted: December 08, 2009
© Santos and Meireles; Licensee Bentham Open.
This is an open access article licensed under the terms of the Creative Commons Attribution Non-Commercial License
(http://creativecommons.org/licenses/by-nc/3.0/) which permits unrestricted, non-commercial use, distribution and reproduction in any medium, provided the
work is properly cited.
218
APÊNDICE IV - MANUAL DE OPERAÇÃO DA UNIDADE MULTIPROPÓSITO
219
220
Manual de Operação da Unidade Multipropósito
ARADIME®
Coordenadora: Profa. Dra. M. Angela A. Meireles
Técnico responsável: Ariovaldo Astini
*Manual elaborado pelo doutorando Diego Tresinari dos Santos
SUMÁRIO
APRESENTAÇÃO ............................................................................................................... 1
FLUXOGRAMA GERAL DA UNIDADE MULTIPROPÓSITO ARADIME® .............. 4
ESPECIFICAÇÕES TÉCNICAS DA UNIDADE MULTIPROPÓSITO ARADIME® .... 5
1A. EXTRAÇÃO UTILIZANDO FLUIDOS SUPERCRÍTICOS (SUPERCRITICAL
FLUID EXTRACTION – SFE) – PARÂMETROS IMPORTANTES NO PROCESSO 6
1B.
HIDRÓLISE
COM
ÁGUA
SUB/SUPERCRÍTICA
COM
CO2
(MODO
BATELADA) – PARÂMETROS IMPORTANTES NO PROCESSO ............................. 8
1.1A SFE sem co-solventes (Procedimento) .................................................................... 10
1.1B Hidrólise com água sub/supercrítica com CO2 (Modo Batelada) (Procedimento) .. 13
FLUXOGRAMA-SFE sem co-solventes/Hidrólise com água sub/supercrítica com
CO2 (Modo Batelada) ................................................................................................... 15
1.2 SFE com co-solventes (Procedimento)....................................................................... 16
FLUXOGRAMA-SFE com co-solventes ..................................................................... 19
1.3 SFE com separadores (Procedimento) ........................................................................ 21
FLUXOGRAMA-SFE com separadores ...................................................................... 24
2A. EXTRAÇÃO COM LÍQUIDOS PRESSURIZADOS (PRESSURIZED LIQUID
EXTRACTION – PLE) – PARÂMETROS IMPORTANTES NO PROCESSO .......... 26
2B. HIDRÓLISE COM ÁGUA SUB/SUPERCRÍTICA SEM CO2 (MODO SEMICONTÍNUO) – PARÂMETROS IMPORTANTES NO PROCESSO .......................... 27
2.1A PLE (Procedimento) ................................................................................................ 28
FLUXOGRAMA-PLE/Hidrólise com água sub/supercrítica sem CO2 (Modo Semicontínuo) ....................................................................................................................... 31
3A. EXTRAÇÃO ASSISTIDA COM DIÓXIDO DE CARBONO A ALTA PRESSÃO
(HIGH PRESSURE CARBON DIOXIDE ASSISTED EXTRACTION-HPCDAE) –
PARÂMETROS IMPORTANTES NO PROCESSO ..................................................... 32
3B. PRÉ-HIDRÓLISE COM EXPLOSÃO COM DIÓXIDO DE CARBONO –
PARÂMETROS IMPORTANTES NO PROCESSO ..................................................... 33
3.1A HPCDAE (Procedimento) ....................................................................................... 35
FLUXOGRAMA-HPCDAE/Pré-tratamento com explosão com dióxido de carbono . 38
ii
4. FORMAÇÃO DE PARTÍCULAS VIA EXPANSÃO RÁPIDA DE SOLUÇÃO
SUPERCRÍTICA (RAPID EXTRACTION OF SUPERCRITICAL SOLUTION –
RESS) – PARÂMETROS IMPORTANTES NO PROCESSO ..................................... 40
4.1 RESS (Procedimento) ................................................................................................. 43
FLUXOGRAMA-RESS/RESS-N ................................................................................ 45
5. FORMAÇÃO DE PARTÍCULAS UTILIZANDO FLUIDO SUPERCRÍTICO COMO
ANTI-SOLVENTE
(SUPERCRITICAL
FLUID
ANTI-SOLVENT
–
SAS)
–
PARÂMETROS IMPORTANTES NO PROCESSO ..................................................... 47
5.1 SAS (Procedimento) ................................................................................................... 50
FLUXOGRAMA-SAS/SAE......................................................................................... 54
ANEXOS ............................................................................................................................. 56
iii
APRESENTAÇÃO
O design de processos industriais sob a óptica da filosofia da
sustentabilidade visa alterações substanciais na indústria atual. Para tanto, é
exigido o desenvolvimento de novos processos baseados em matérias-primas
renováveis, na utilização mínima necessária de energia e solventes sem restrições
ambientais. Neste contexto, as tecnologias baseadas na utilização de fluidos
pressurizados parecem oferecer soluções para essas demandas.
Tecnologias de extração de compostos bioativos de vegetais podem
representar uma alternativa ambientalmente correta e economicamente viável aos
métodos convencionais de extração, onde grandes quantidades de solventes,
longos tempos de extração e altas temperaturas são requeridas, o que pode
adicionalmente promover a degradação destes compostos durante o processo
extrativo.
O LASEFI (LAboratório de tecnologia Supercrítica: Extração, Fracionamento
e Identificação de extratos vegetais) em atividade desde 1984 possui uma vasta
experiência no estudo de processos para a produção de extratos de plantas
aromáticas, condimentares e medicinais, assim como, o fracionamento de extratos
para obtenção de frações concentradas bioativas através do uso de tecnologias de
produção limpas, que preservem o meio ambiente e não tragam danos à saúde
humana e animal, em todas as etapas de processamento, através do uso de
tecnologias inovadoras, como a extração e o fracionamento com fluídos
pressurizados. Outro foco do LASEFI também é o design e montagem de
extratores supercríticos. O LASEFI possui em suas instalações dois extratores
supercríticos construídos: um menor que opera com CO2 supercrítico com ou sem
co-solventes e outro maior (células extratoras de 1 L) que opera somente com
CO2 supercrítico, porém possibilita o reciclo do solvente, bem como a operação
em modo contínuo, pois possui duas células extratoras em paralelo, além de
outros dois comerciais (Applied Separations e Thar) e um construído pela equipe
do Professor Brunner.
Utilizando todo o conhecimento construído pelo grupo de pesquisa da
Profa. Dra. Maria Angela de Almeida Meireles visando à extensão da área de
1
atuação do LASEFI uma nova unidade para o desenvolvimento de processos que
utilizam fluidos pressurizados foi projetada, montada e testada com sucesso.
Batizada de ARADIME®, devido ela ter sido elaborada pelos profissionais
ARiovaldo Asitini, DIego Tresinari dos Santos e Maria Angela de Almeida MEireles
a inovadora unidade além de possibilitar o desenvolvimento de extrações
supercríticas utilizando CO2 supercrítico com ou sem co-solventes, ou somente
utilizando solventes líquidos pressurizados, proporciona a possibilidade de
desenvolver extrações assistidas com dióxido de carbono a alta pressão,
formação de partículas encapsuladas ou não, reação utilizando fluidos
supercríticos, além de outros processos.
Técnicas de formação de partículas utilizando fluidos pressurizados acima
da condição supercrítica (conhecidos como fluidos supercríticos) podem: i) permitir
um aumento no poder de dissolução de compostos através da redução do
tamanho das partículas; ii) proteção/estabilidade de compostos sem levar o aditivo
às condições que podem ocasionar sua degradação, isto é, perda de cor,
capacidade antioxidante, etc, condições estas normalmente utilizadas nos
processos convencionais. Adicionalmente, processos de precipitação utilizando
fluidos supercríticos permitem um fácil controle da formação de partículas através
de pequenas variações nas condições de operação (Pressão, Temperatura, etc).
Diferentes processos de micronização/encapsulação que usam fluidos
supercríticos, bem como equipamentos para a realização destes processos têm
sido desenvolvidos. Estes processos podem ser classificados de acordo com a
função do fluido supercrítico no processo: solvente [“Rapid Expansion of
Supercritical Solutions” (RESS)]; “Supercritical Solvent Impregnation” (SSI); soluto
[“Particles from Gas Saturated Solutions” (PGSS)] ou anti-solvente [“Supercritical
Anti-Solvent” (SAS); “Supercritical Fluid Extraction of Emulsions” (SFEE).
A seguir são descritos os procedimentos de operação da unidade
ARADIME® para o desenvolvimento de alguns dos diversos processos que ela
pode operar: Processos de extração utilizando Fluidos Supercríticos (Supercritical
Fluid Extraction – SFE) com ou sem a adição de co-solventes, com ou sem a
utilização de separadores; extração com líquidos pressurizados (Pressurized
2
Liquid Extraction – PLE); extração assistida com dióxido de carbono a alta pressão
(High Pressure Carbon Dioxide Assisted Extraction-HPCDAE); pré-hidrólise com
explosão com CO2 supercrítico; hidrólise com água subcrítica com ou sem adição
de CO2, de modo semi-contínuo ou em batelada; formação de partículas via
expansão rápida de solução supercrítica (Rapid Extraction of Supercritical Solution
– RESS) e formação de partículas utilizando fluidos supercrítico como antisolvente (Supercritical fluid Anti-Solvent – SAS).
Para facilitar o entendimento sobre os processos que a unidade pode
desenvolver, um resumo sobre a influência dos parâmetros mais importantes em
cada processo é apresentado. Já para facilitar o entendimento de como a unidade
multipropósito funciona um fluxograma geral é apresentado a seguir. Neste
fluxograma todos os componentes que compõem a unidade estão desenhados.
Percebe-se que parte dos componentes estão conectadas a outras, e parte não.
As partes que não estão conectadas (exemplo: BL, V-5, V-6, V-7, etc.) foram
desenhadas assim, pois elas podem ser conectadas de diferentes formas para
possibilitar o desenvolvimento de diferentes processos, através do emprego de
alguns acessórios. Já as partes que estão conectadas (exemplo: V-1, FL, M-1, VC,
etc.) sempre vão permanecer como estão para o desenvolvimento de todos os
processos.
Nos fluxogramas dos processos que unidade desenvolve, foi-se adotado
que as partes não conectadas serão conectadas aos seus devidos lugares para
possibilitar o desenvolvimento do processo desejado através da utilização de uma
linha tracejada espessa. Adicionalmente, uma lista dos componentes que não
serão utilizados durante o desenvolvimento de cada processo é mencionada para
evitar equivocações por parte do operador, bem como imagens de como a
unidade deve estar para desenvolver cada um dos processos são apresentadas.
Ao final deste manual (Seção ANEXOS), encontram-se tabelas contendo
valores da densidade do CO2 em condições sub/supercríticas e em condições
encontradas no laboratório, valores da viscosidade da água em condições
sub/supercríticas e valores das constantes dielétricas de alguns solventes
orgânicos.
3
FLUXOGRAMA GERAL DA UNIDADE MULTIPROPÓSITO ARADIME®
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
BG
V-7
V-BPR
V-AR
M-3
VS
BR
V-3
V-5
V-4
V-BR-1
VP-1
V-BR-2
VP-2
BL
BA
FL – Filtro de Linha; V-1, V-2, V-3, V-4, V-5, V-6, V-7
– Válvulas de Bloqueio; V-BPR – Válvula Back Pressure Regulator; VC –
Válvula Compressor (Controle da Vazão de ar Comprimido); V-AR – Válvula Anti-Retorno; V-BR-1, V-BR-2 – Válvulas Banho de
Refrigeração (Controle da Circulação do Fluido Refrigerante); VM – Válvula micrométrica; VS – Válvula de Segurança; M-1, M-2, M3, M-4 – Manômetros; BR – Banho de Resfriamento; BA – Banho de Aquecimento; BG – Bomba para Gases (em condições
atmosféricas); BL – Bomba para Líquidos (em condições atmosféricas); VP-1 – Vaso de Pressão pequeno (6,57 mL); VP-2 – Vaso de
Pressão Grande (500 mL); CAVP-1 – Controlador do Aquecimento do Vaso de Pressão 1; CAVM – Controlador do Aquecimento da
Válvula Micrométrica; TP – Termopar; RT – Rotâmetro; TO – Totalizador.
4
ESPECIFICAÇÕES TÉCNICAS DA UNIDADE MULTIPROPÓSITO ARADIME®
Faixa de Temperatura: Vaso de Pressão Pequeno (6,57 mL) – Ambiente a 400 ºC;
Vaso de Pressão Grande (500 mL) - -10ºC a 250 ºC; Válvula Micrométrica –
Temperatura limite de 120 ºC;
Faixa de Pressão: Vaso de Pressão Pequeno (6,57 mL) – Pressão limite de 450
bar; Vaso de Pressão Grande (500 mL) - Pressão limite de 200 bar;
Faixa de Vazão: CO2 - 0.15-2.2 kg/h de CO2 a 1.013bar/20 ºC; Líquido – 0.01-9,99
mL/min.
Acessórios:
1) Tubulação VP-1/Entrada
2) Tubulação VP-1/Saída
3) Tubulação VM/1
4) Tubulação VM/2
5) Tubulação VP-2
6) Tubulação V-7
7) Tubulação RT
8) Tubulação T
Diâmetro interno do capilar interno do sistema coaxial (Tubulação T): 177,8 μm
Diâmetro interno do bico expansor (Tubulação VM/2): 0,51 mm
5
1a. Extração utilizando Fluidos Supercríticos (Supercritical Fluid Extraction – SFE)
– Parâmetros importantes no Processo
-
Pressão;
-
Temperatura;
-
Vazão do solvente (mais comum é o CO2);
-
Vazão do co-solvente - Relação entre co-solvente/solvente (co-solventes
mais aconselháveis são: etanol, isopropanol e água).
A variação na Pressão e Temperatura de extração está diretamente relacionada a
variação na densidade do fluido supercrítico. Sendo a seletividade do processo
extrativo ajustada através desta mudança.
Segundo a experiência do LASEFI, o comportamento do rendimento de extração
(massa de extrato/massa de matéria-prima) se assemelha ao da solubilidade de
solutos em fluido supercrítico. Portanto, o método dinâmico de determinação de
solubilidade dos solutos a serem extraídos pode ser aplicado eficazmente
utilizando o mesmo equipamento de extração supercrítica. Tal método avalia a
massa de extrato obtida (solutos solubilizados) ao se empregar diferentes
pressões e temperaturas. Geralmente 4 ou 5 pressões e duas temperaturas são
utilizadas.
6
Para processos de extração e posterior fracionamento dos extratos os parâmetros
pressão e temperatura dos vasos separadores, bem como a vazão do fluido
supercrítico continuam sendo importantes. O fracionamento dos extratos nos
separadores se dá através da modificação da solubilidade dos solutos-alvo,
variando os parâmetros dos processos mencionados. Outra forma de se obter
extratos fracionados é desenvolver extrações de uma mesma matéria-prima em
várias etapas, empregando diferentes solventes pressurizados. O caso mais
comum envolve o emprego de CO2 supercrítico puro primeiramente (obtendo um
extrato rico em compostos mais apolares), seguido de um posterior emprego de
CO2 + etanol (com um teor máximo de etanol de 20% m/m) (obtendo um extrato
rico em compostos mais polares).
A extração com fluidos supercríticos utilizando altos teores de co-solventes (mais
que 20 % m/m) é conhecida como extração acelerada com solvente, extração
acelerada por solvente ou extração por solvente acelerado (Accelerated Solvent
Extraction (ASE)), uma vez que uma alta concentração de co-solvente (solvente
líquido) acelera o procedimento de extração. Tal método extrativo tem se
demonstrado promissor para a extração de compostos mais polares como
curcuminóides, flavonóides, etc. Mais abrangentemente alguns grupos de
pesquisa chamam de “Enhanced Solvent Extraction (ESE)” a extração que utiliza
7
fluidos supercríticos em combinação com um ou mais solventes em quaisquer
proporções.
Quando CO2 e água são conjuntamente empregados nas extrações se é
observado um decréscimo no pH através da geração in situ de ácido carbônico, o
que leva a uma melhora na extração de alguns compostos, tais como
antocianinas, por exemplo. Dependendo da temperatura empregada (> 150 ºC)
concomitantemente com a extração, CO2 e água podem atuar como meio
reacional proporcionando uma indesejável hidrólise da matriz vegetal.
1b. Hidrólise com água sub/supercrítica com CO2 (Modo Batelada) – Parâmetros
importantes no Processo
-
Pressão;
-
Temperatura;
-
Relação massa de matéria-prima/massa de água na alimentação do reator;
-
Tempo de reação;
-
Vazão de saída do CO2.
Água a altas temperaturas e pressões apresenta propriedades únicas, como a
possibilidade de modificar o produto iônico, a constante dielétrica e a densidade, o
8
que a torna um meio de reação interessante. A água subcrítica apresenta
temperaturas entre 150 e 370 °C e pressões entre 4 e 22 MPa. Nestas condições,
a água pode agir como um catalisador ácido, acelerando o processo de hidrólise.
Valores de temperatura e pressão acima de 374 °C/22 MPa caracterizam água
supercrítica. A fim de acidificar o meio aquoso sub/supercrítico para acelerar o
processo de hidrólise conforme mencionado anteriormente, dióxido de carbono
(CO2) pode ser adicionar ao meio. O CO2 reage com a água formando ácido
carbônico, o qual age como catalisador do processo.
Palha de trigo, folhas, caule e espiga de milho, bagaço de gengibre, farelo de
arroz, bagaço de cana de açúcar, entre outros, têm sido eficazmente hidrolisados
utilizando água sub/supercrítica com ou sem CO2 visando a obtenção de produtos
de alto valor agregado, tais como etanol, xilitol, sorbitol, vitamina C, polímeros, etc.
9
1.1A SFE sem co-solventes (Procedimento)
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Inserir a célula de extração (VP-1) contendo a matéria-prima rica em compostos
bioativos a serem extraídos (geralmente, já devidamente seca e moída) no
sistema de aquecimento do VP-1 desligado;
3) Conectar a tubulação VP-1/Entrada entre a válvula V-5 e o topo do VP-1;
4) Conectar a tubulação VP-1/Saída entre o fundo do VP-1 e a válvula V-6;
5) Conectar a tubulação VM/1 na tampa de um frasco coletor lacrado (50 ou 100
mL) de massa conhecida, imerso em um banho de gelo;
6) Conectar a tubulação RT também na tampa do frasco coletor;
7) Verificar se as válvulas V-1, V-2, V-5, V-6 e VM estão fechadas;
8) Abrir a válvula do cilindro de CO2 e a válvula V-1 lentamente, observando se o
ponteiro do manômetro M-1 sobe suavemente;
9) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
10) Quando a pressão do CO2 for atingida, abrir a válvula V-5 para pressurizar a
célula extratora (VP-1);
11) Ligar o controlador do sistema de aquecimento do vaso de pressão 1 (CAVP1) e da válvula micrométrica (CAVM) e programá-los para operar à temperatura
10
requerida do processo e a uma temperatura que evite congelamento da linha de
saída, respectivamente (esta temperatura depende da vazão de CO2 utilizada,
podendo oscilar entre 70 a 120 ºC);
12) Quando a pressão e temperatura do sistema estiverem estabilizadas,
cronometrar o tempo estipulado para o período estático (Este tempo, quando
requerido, pode oscilar de 5 a 30 minutos);
13) Anotar o volume indicado no totalizador (TO);
14) Após o tempo do período estático, abrir as válvulas V-6 e VM cuidadosamente
até que a vazão desejada seja alcançada (Medição através do rotâmetro);
15) Após o tempo pré-estabelecido de extração, desligar o banho BR e os
controladores de aquecimentos CAVP-1 e CAVM e fechar as válvulas V-5, V-1 e
VC;
16) Anotar o volume indicado no totalizador (TO) e contabilizar o volume de CO2
que efetivamente foi percolado durante a extração;
17) Despressurizar o vaso VP-1;
18) Limpar a tubulação de saída com um solvente (etanol, por exemplo);
19) Quantificar o teor de extrato residual na tubulação de saída através da
evaporação do solvente e pesagem da massa de extrato seca;
20) Quantificar a massa de extrato obtida através da pesagem do frasco coletor
(Geralmente, a massa de extrato residual na tubulação de saída é somada a
massa de extrato obtida para fins de determinação de rendimento);
11
21) Guardar o frasco coletor em freezer à temperatura igual ou menor que zero
para posterior análise.
12
1.1B
Hidrólise
com
água
sub/supercrítica
com
CO2
(Modo
Batelada)
(Procedimento)
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Inserir o reator de hidrólise (VP-1) contendo a biomassa a ser hidrolisada com
uma determinada quantidade de água ou umidade intrínseca no sistema de
aquecimento do VP-1 desligado;
3) Conectar a tubulação VP-1/Entrada entre a válvula V-5 e o topo do VP-1;
4) Conectar a tubulação VP-1/Saída entre o fundo do VP-1 e a válvula V-6;
5) Conectar a tubulação VM/1 na tampa de um frasco coletor lacrado (50 ou 100
mL) de massa conhecida, imerso em um banho de gelo;
6) Conectar a tubulação RT também na tampa do frasco coletor;
7) Verificar se as válvulas V-1, V-2, V-5, V-6 e VM estão fechadas;
8) Abrir a válvula do cilindro de CO2 e a válvula V-1 lentamente, observando se o
ponteiro do manômetro M-1 sobe suavemente;
9) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
10) Ligar o controlador do sistema de aquecimento do vaso de pressão 1 (CAVP1) e programá-lo para operar à temperatura requerida do processo;
13
11) Quando a pressão do CO2 e a temperatura do vaso VP-1 for atingida, abrir a
válvula V-5 para pressurizar o reator de hidrólise (VP-1);
12) Quando a pressão e temperatura do sistema estiverem estabilizadas,
cronometrar o tempo estipulado para o tempo de reação e fechar a válvula V-5;
13) Decorrido o tempo de hidrólise, abrir as válvulas V-6 e VM cuidadosamente
para que os produtos de reação sejam arrastados do reator na vazão desejada
(Medição através do rotâmetro);
14) Após a eliminação de todo o hidrolisado, desligar o banho BR e os
controladores de aquecimentos CAVP-1 e CAVM e fechar as válvulas V-1 e VC;
15) Utilizar um ventilador para ajudar a resfriar o vaso VP-1 mais rapidamente;
16) Pesar a biomassa residual contida dentro do reator (VP-1);
17) Quantificar a massa de hidrolisado obtida através da pesagem do frasco
coletor (Geralmente, a massa de hidrolisado obtido somado a massa da biomassa
residual dá valores próximos à massa de biomassa inicial; caso isto não ocorra o
experimento deve ser repetido);
18) Guardar o frasco coletor e o material residual em freezer à temperatura igual
ou menor que zero para posterior análise.
14
FLUXOGRAMA-SFE sem co-solventes/Hidrólise com água sub/supercrítica com CO2 (Modo Batelada)
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
BG
V-7
V-BPR
V-AR
M-3
BR
V-3
VS
V-5
V-4
V-BR-1
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo SFE sem co-solventes/Hidrólise com água
sub/supercrítica com CO2 (Modo Batelada)
V-7
– Válvula de Bloqueio; V-BR-1, V-BR-2 – Válvulas Banho de Refrigeração (Controle da Circulação do Fluido
Refrigerante); M-4 – Manômetro; BA – Banho de Aquecimento; BL – Bomba para Líquidos (em condições atmosféricas);
VP-2 – Vaso de Pressão Grande (500 mL); Termopar.
15
1.2 SFE com co-solventes (Procedimento)
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Inserir a célula de extração (VP-1) contendo a matéria-prima rica em compostos
bioativos a serem extraídos (geralmente, já devidamente seca e moída) no
sistema de aquecimento do VP-1 desligado;
3) Conectar a bomba para Gases à válvula V-4;
4) Conectar a tubulação VP-1/Entrada entre a válvula V-5 e o topo do VP-1;
5) Conectar a tubulação VP-1/Saída entre o fundo do VP-1 e a válvula V-6;
6) Conectar a tubulação VM/1 na tampa de um frasco coletor lacrado (50 ou 100
mL) de massa conhecida, imerso em um banho de gelo;
7) Conectar a tubulação RT também na tampa do frasco coletor;
8) Verificar se as válvulas V-1, V-2, V-4, V-5, V-6 e VM estão fechadas;
9) Abrir a válvula do cilindro de CO2 e a válvula V-1 lentamente, observando se o
ponteiro do manômetro M-1 sobe suavemente;
10) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
11) Ligar a bomba para líquidos (BL) e programá-la para operar à vazão requerida
do processo;
16
12) Ligar o controlador do sistema de aquecimento do vaso de pressão 1 (CAVP1) e da válvula micrométrica (CAVM) e programá-los para operar à temperatura
requerida do processo e a uma temperatura que evite congelamento da linha de
saída, respectivamente (esta temperatura depende da vazão de CO2 utilizada,
podendo oscilar entre 70 a 120 ºC);
13) Anotar o volume indicado no totalizador (TO);
14) Quando a pressão do CO2 for atingida, simultaneamente, abrir a válvula V-4 e
V-5 para pressurizar a célula extratora (VP-1) com a mistura CO2 + co-solvente;
15) Após a pressão e temperatura do sistema estiverem estabilizadas, abrir as
válvulas V-6 e VM cuidadosamente até que a vazão do CO2 desejada seja
alcançada (Medição através do rotâmetro) (Se assume que a vazão de cosolvente programada permanecerá sempre constante durante o processo de
extração);
16) Após o tempo pré-estabelecido de extração, desligar o banho BR, os
controladores de aquecimentos CAVP-1 e CAVM e a bomba BL e fechar as
válvulas V-5, V-1 e VC;
17) Anotar o volume indicado no totalizador (TO) e contabilizar o volume de CO2
que efetivamente foi percolado durante a extração;
18) Despressurizar o vaso VP-1;
19) Limpar a tubulação de saída com um solvente (etanol, por exemplo);
17
20) Quantificar o teor de extrato residual na tubulação de saída através da
evaporação do solvente e pesagem da massa de extrato seca;
21) Quantificar a massa de extrato obtida através da evaporação do solvente e
pesagem da massa de extrato seca (Geralmente, a massa de extrato residual na
tubulação de saída é somada a massa de extrato obtida para fins de determinação
de rendimento);
22) Guardar o frasco coletor em freezer à temperatura igual ou menor que zero
para posterior análise.
18
FLUXOGRAMA-SFE com co-solventes
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
BG
V-7
V-BPR
V-AR
M-3
BR
V-3
VS
V-5
V-4
V-BR-1
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo SFE com co-solventes
V-7
– Válvula de Bloqueio; V-BR-1, V-BR-2 – Válvulas Banho de Refrigeração (Controle da Circulação do Fluido
Refrigerante); M-4 – Manômetro; BA – Banho de Aquecimento; VP-2 – Vaso de Pressão Grande (500 mL); Termopar.
19
20
1.3 SFE com separadores (Procedimento)
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Se a temperatura do vaso separador desejada for: i) -10 ºC, abrir as válvulas
banho de refrigeração (V-BR-1 e V-BR-2); ii) maior do que a temperatura
ambiente, ligar o banho de aquecimento (BA) e programá-lo para operar na
temperatura desejada; iii) entre -10 ºC e a temperatura ambiente, abrir as válvulas
banho de refrigeração (V-BR-1 e V-BR-2), ligar o banho de aquecimento (BA) e
programá-lo para operar em uma temperatura que possibilite atingir a temperatura
desejada do vaso separador;
3) Inserir a célula de extração (VP-1) contendo a matéria-prima rica em compostos
bioativos a serem extraídos (geralmente, já devidamente seca e moída) no
sistema de aquecimento do VP-1 desligado;
4) Conectar a tubulação VP-1/Entrada entre a válvula V-5 e o topo do VP-1;
5) Conectar a tubulação VP-1/Saída entre o fundo do VP-1 e a válvula V-6;
6) Conectar a tubulação VP-2 entre o orifício lateral (esquerda) do vaso separador
(VP-2) e a válvula V-7;
7) Conectar a tubulação V-7 na tampa de um frasco coletor lacrado (50 ou 100
mL) de massa conhecida, imerso em um banho de gelo;
8) Conectar a tubulação RT também na tampa do frasco coletor;
21
9) Conectar a tubulação VM/2 no orifício central do vaso separador (VP-2);
10) Verificar se as válvulas V-1, V-2, V-5, V-6, VM e V-7 estão fechadas;
11) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
12) Ligar o controlador do sistema de aquecimento do vaso de pressão 1 (CAVP1) e da válvula micrométrica (CAVM) e programá-los para operar à temperatura
requerida do processo e a uma temperatura que evite congelamento da linha de
saída, respectivamente (esta temperatura depende da vazão de CO2 utilizada,
podendo oscilar entre 70 a 120 ºC);
13) Lacrar o vaso separador (VP-2), conectando o termopar e o manômetro M-4
ao vaso VP-2;
14) Quando a pressão do CO2 for atingida, abrir a válvula V-5 para pressurizar a
célula extratora (VP-1);
15) Após a pressão e temperatura da célula extratora estiverem estabilizadas,
abrir as válvulas V-6 e VM até que o CO2 preencha o volume do vaso separador
(VP-2);
16) Anotar o volume indicado no totalizador (TO);
17) Uma vez que a pressão do vaso separador esteja próxima a desejada
(Medição através do manômetro M-4), abrir a válvula V-7 cuidadosamente e
controlar a válvula VM até que a vazão do CO2 desejada seja alcançada (Medição
22
através do rotâmetro) e a pressão do vaso separador (VP-2) permaneça
constante;
18) Após o tempo pré-estabelecido de extração, desligar os banho BR e BA, os
controladores de aquecimentos CAVP-1 e CAVM e a bomba BL e fechar as
válvulas V-5, V-1 e VC;
19) Anotar o volume indicado no totalizador (TO) e contabilizar o volume de CO2
que efetivamente foi percolado durante a extração;
20) Despressurizar os vasos VP-1 e VP-2;
21) Quantificar a massa de extrato obtida no vaso separador (VP-2) e no frasco
coletor, através da coleta do extrato precipitado no vaso VP-2 e da pesagem do
frasco coletor, respectivamente;
22) Guardar ambos os extratos em freezer à temperatura igual ou menor que zero
para posterior análise.
OBS: Para o desenvolvimento de extrações SFE com separadores e co-solventes,
verificar na seção 1.2 que algumas etapas são diferentes, como a evaporação do
solvente ao final da extração para se quantificar a massa de extrato obtida, por
exemplo.
23
FLUXOGRAMA-SFE com separadores
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
BG
V-7
V-BPR
V-AR
M-3
BR
V-3
VS
V-5
V-4
V-BR-1
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo SFE com co-solventes
Todos os componentes são utilizados
24
25
2a. Extração com Líquidos Pressurizados (Pressurized Liquid Extraction – PLE) –
Parâmetros importantes no Processo
-
Temperatura;
-
Vazão do líquido pressurizado;
-
Tempo da extração estática.
A variação na Temperatura de extração a uma Pressão relativamente baixa (50100 bar) está diretamente relacionada a variação na viscosidade do líquido
pressurizado, que permanece no estado líquido bem acima da sua temperatura de
ebulição. Sendo a eficiência do processo extrativo ajustada através desta
mudança.
Segundo a literatura e a experiência do LASEFI no processo PLE um aumento na
pressão de extração não afeta significamente o processo extrativo. Portanto,
geralmente prefere-se se trabalhar com pressões baixas (50-70 bar).
Uma atenção redobrada deve ser dada a vazão do líquido pressurizado e o tempo
da extração estática para a extração de compostos termosensíveis, uma vez que
elas podem resultar em um tempo excessivamente grande que pode iniciar a
degradação destes compostos durante o procedimento de extração.
26
A extração com líquidos pressurizados também é chamada por alguns autores de
extração acelerada com solvente, como a extração supercrítica com altos teores
de co-solvente, pelos mesmos motivos, isto é, propiciar uma extração rápida dos
solutos.
2b. Hidrólise com água sub/supercrítica sem CO2 (Modo Semi-contínuo) –
Parâmetros importantes no Processo
-
Pressão;
-
Temperatura;
-
Tempo de reação;
-
Relação massa de matéria-prima/água percolada;
-
Vazão de alimentação da água sub/supercrítica.
Conforme já mencionado anteriormente, água a altas temperaturas (> 150 ºC) e
pressões é considerada um meio reacional muito atrativo para reações de
hidrólise.
27
2.1A PLE (Procedimento)
1) Inserir a célula de extração (VP-1) contendo a matéria-prima rica em compostos
bioativos a serem extraídos [geralmente, cortada em pedaços pequenos com ou
sem a adição de um suporte inerte (areia, celite, etc.)] no sistema de aquecimento
do VP-1 desligado;
2) Conectar a tubulação VP-1/Entrada entre a válvula V-5 e o topo do VP-1;
3) Conectar a tubulação VP-1/Saída entre o fundo do VP-1 e a válvula V-6;
4) Conectar a tubulação VM/1 em um frasco coletor 25 ou 50 mL de massa
conhecida, imerso em um banho de gelo;
5) Verificar se as válvulas V-4, V-5, V-6 e VM estão fechadas;
6) Ligar a bomba para líquidos (BL) e programá-la para operar à pressão
requerida do processo [uma oscilação entre o valor da pressão normalmente é
observada durante o processo extrativo, portanto a bomba deve ser programada
para operar a uma pressão um pouco maior (10-15 bar maior)];
7) Ligar o controlador do sistema de aquecimento do vaso de pressão 1 (CAVP-1)
e programá-lo para operar à temperatura requerida do processo;
8) Abrir a válvula V-4, verificando o aumento da pressão (Medição através do
manômetro M-3);
9) Quando a pressão do líquido for atingida, abrir a válvula V-5 para pressurizar a
célula extratora (VP-1);
28
10) Quando a pressão e temperatura do sistema estiverem estabilizadas,
cronometrar o tempo estipulado para o período estático (Este tempo pode oscilar
de 3 a 15 minutos);
11) Após o tempo do período estático, abrir as válvulas V-6 e VM, muito
cuidadosamente, (pois, a pressão pode cair drasticamente) até que a vazão
desejada seja alcançada (Medição utilizando proveta e cronômetro);
12) Após o tempo pré-estabelecido de extração, desligar a bomba BL e o
controlador de aquecimento CAVP-1 e fechar a válvula V-4;
13) Para eliminar o extrato residual-solvente que ficou na célula extratora, purgar
CO2 através da abertura das válvulas: do cilindro, V-1 e V-2.
14) Quantificar o teor de extrato residual na célula extratora através da
evaporação do solvente (o qual foi purgado pelo CO2) e pesagem da massa de
extrato seca;
15) Quantificar a massa de extrato obtida através da evaporação do solvente e
pesagem da massa de extrato seca (Geralmente, a massa de extrato residual na
célula extratora é somada a massa de extrato obtida para fins de determinação de
rendimento);
16) Guardar o frasco coletor em freezer à temperatura igual ou menor que zero
para posterior análise.
29
OBS 1: Para o desenvolvimento de extrações PLE com água como solvente, o
procedimento de quantificação da massa do extrato fica mais complexo. Algumas
opções são: através de secagem em chapa aquecida, de secagem com nitrogênio
a quente, de liofilização do extrato aquoso, entre outras. Em contrapartida, as
análises
químicas
do
extrato
aquoso
podem
ser
feitas
normalmente
(Determinação da Atividade Antioxidante, Determinação do Teor de Compostos
Fenólicos, etc.).
OBS 2: Para o desenvolvimento de processos de hidrólise com água
sub/supercrítica sem CO2 (Modo Semi-contínuo), seguir procedimento exatamente
igual ao descrito, diferenciando que nunca se deve adicionar um suporte inerte
juntamente à biomassa a ser hidrolisada. Diferentemente do processo descrito na
seção 1.1b este processo não ocorre em modo batelada e sim em modo semicontínuo, portanto o meio reacional não é alimentado no início do processo e sim
continuamente, até o término da reação. “Lembrar que para se conseguir obter
hidrolisados a temperatura que a água deve estar deve ser maior que 150 ºC”.
30
FLUXOGRAMA-PLE/Hidrólise com água sub/supercrítica sem CO2 (Modo Semi-contínuo)
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
BG
V-7
V-BPR
V-AR
M-3
BR
V-3
VS
V-5
V-4
V-BR-1
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo PLE/Hidrólise com água sub/supercrítica sem CO2
(Modo Semi-contínuo)
V-7
– Válvula de Bloqueio; VC – Válvula Compressor (Controle da Vazão de ar Comprimido); V-BR-1, V-BR-2 – Válvulas
Banho de Refrigeração (Controle da Circulação do Fluido Refrigerante); M-4 – Manômetro; BA – Banho de Aquecimento; BG –
Bomba para Gases (em condições atmosféricas); VP-2 – Vaso de Pressão Grande (500 mL); CAVM – Controlador do
Aquecimento da Válvula Micrométrica; TP – Termopar; RT – Rotâmetro; TO – Totalizador.
31
3a. Extração assistida com dióxido de carbono a alta pressão (High Pressure
Carbon Dioxide Assisted Extraction-HPCDAE) – Parâmetros importantes no
Processo
-
Pressão;
-
Temperatura;
-
Tempo da extração;
-
Relação massa de matéria-prima/volume de solvente adicionado ao vaso
de pressão;
-
Relação volume de CO2/volume do solvente+matéria-prima no vaso de
pressão;
-
Velocidade da etapa de despressurização.
Devido à similaridade entre os processos que visam à inativação microbiana e os
que visam à extração sólido-líquido de compostos bioativos de matrizes vegetais,
alguns autores deduziram e confirmaram com seus resultados experimentais que
extrações sólido-líquido assistidas com CO2 a alta pressão poderiam propiciar
melhores resultados que quando não assistidas.
O efeito explosivo do CO2 a alta pressão em primeiro lugar foi demonstrado
romper células bacterianas através da rápida liberação de pressão de gás com o
objetivo de recolher o conteúdo da célula (lise celular). Numerosos estudos têm
32
mostrado a eficácia do uso de CO2 a alta pressão para inativar microrganismos e
enzimas
A melhora do processo de extração através do uso de CO2 de alta pressão é
atribuída às habilidades do CO2 a alta pressão modificar a membrana celular,
diminuir o pH intracelular, desorientar o equilíbrio eletrolítico intracelular, remover
os componentes vitais das células e membranas celulares.
Quando o solvente empregado nas extrações é a água, o uso de CO2 a alta
pressão possibilita um decréscimo no pH através da geração in situ de ácido
carbônico, o que leva a uma melhora na extração de alguns compostos, tais como
antocianinas, por exemplo.
3b. Pré-hidrólise com explosão com dióxido de carbono – Parâmetros importantes
no Processo
-
Pressão;
-
Temperatura;
-
Tempo do pré-tratamento;
-
Relação volume de CO2/volume da biomassa no vaso de pressão;
-
Umidade da biomassa;
-
Velocidade da etapa de despressurização.
33
Diversas
técnicas
vêm
sendo
empregadas
objetivando-se
aumentar
a
digestibilidade de biomassas (facilitar a hidrólise).
O pré-tratamento utilizando explosão com CO2 nas condições supercríticas surge
como uma alternativa ao método convencional de explosão com vapor. O
processo consiste em submeter às amostras nas condições supercríticas por um
tempo determinado, de forma a permitir que o CO2 penetre nas estruturas do
material, ocupando assim os espaços vazios. Dessa forma após a rápida
despressurização, o CO2 retornaria na forma de gás provocando assim uma
alteração na estrutura celular.
A umidade da biomassa a ser pré-tratada tem demonstrado exercer importante
efeito sobre o material a ser hidrolisado. Visando a posterior aplicação da
biomassa para outros objetivos que não a hidrólise, tais como, remoção de metais
pesados, imobilização de células microbianas, entre outros, um tratamento mais
brando é requerido, isto é utilizando uma biomassa com baixo conteúdo de água.
34
3.1A HPCDAE (Procedimento)
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Conectar a tubulação VP-2 entre a válvula V-5 e o orifício lateral (esquerda) do
vaso separador (VP-2);
3) Conectar a tubulação T no orifício central do vaso separador (VP-2) e suas
ramificações na válvula V-6 e na bomba BL (a conexão na bomba BL serve
somente para estancar o CO2);
4) Verificar se as válvulas V-1, V-2, V-5 e V-6 estão fechadas e se a VM está
aberta;
5) Abrir a válvula do cilindro de CO2 e a válvula V-1 lentamente, observando se o
ponteiro do manômetro M-1 sobe suavemente;
6) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
7) Ligar o controlador do sistema de aquecimento da válvula micrométrica (CAVM)
e programá-lo para operar à temperatura que evite congelamento da linha de
saída, respectivamente (geralmente, esta temperatura deve estar ao redor de 100120 ºC, pois a descompressão do CO2 neste processo deve ser realizada de
forma muito rápida);
35
8) Ligar o banho de aquecimento (BA) e programá-lo para operar na temperatura
desejada;
9) Inserir na célula de extração (VP-2) o solvente de extração a ser utilizado e a
matéria-prima rica em compostos bioativos a serem extraídos (Diferentes
Relações massa de matéria-prima/volume de solvente adicionado ao vaso de
pressão e Relações volume de CO2/volume do solvente+matéria-prima no vaso de
pressão podem ser avaliadas);
10) Lacrar a célula de extração (VP-2), conectando o termopar e o manômetro M-4
ao vaso VP-2;
11) Quando a pressão e temperatura do sistema estiverem estabilizadas,
cronometrar o tempo estipulado para a extração e fechar a válvula V-6 (Este
tempo pode oscilar de 5 a 50 minutos);
12) Decorrido o tempo de extração, abrir a válvula V-6 rapidamente,
despressurizando o vaso VP-2 (O tempo de despressurização deve ser menor do
que 10 minutos);
13) Deslacrar a célula de extração (VP-2), coletar o extrato (aquoso, etanólico, etc)
e o refrigerar através da imersão em banho de gelo (para evitar degradação);
14) Desligar os banhos BR e BA e o controlador de aquecimento CAVM e fechar
as válvulas V-1 e VC;
36
15) Quantificar a massa de extrato obtida através da evaporação do solvente e
pesagem da massa de extrato seca;
16) Guardar o extrato em freezer à temperatura igual ou menor que zero para
posterior análise.
OBS 1: Para o desenvolvimento de extrações PLE com água como solvente, o
procedimento de quantificação da massa do extrato fica mais complexo. Algumas
opções são: através de secagem em chapa aquecida, de secagem com nitrogênio
a quente, de liofilização do extrato aquoso, entre outras. Em contrapartida, as
análises
químicas
do
extrato
aquoso
podem
ser
feitas
normalmente
(Determinação da Atividade Antioxidante, Determinação do Teor de Compostos
Fenólicos, etc.).
OBS 2: Para o desenvolvimento de processos de pré-hidrólise com explosão com
CO2 supercrítico, seguir procedimento exatamente igual ao descrito, diferenciando
que nenhum solvente deve ser adicionado juntamente à biomassa a ser
hidrolisada. Em alguns casos onde se é desejado uma pré-hidrólise mais efetiva, a
umidade da biomassa é aumentada através da impregnação de uma determinada
massa de água à biomassa.
37
FLUXOGRAMA-HPCDAE/Pré-tratamento com explosão com dióxido de carbono
CAVM
TP
M-4
M-2
VC
M-1
CAVP-1
TO
RT
V-6
VM
V-2
V-1
FL
V-7
BG
V-BPR
V-AR
M-3
BR
V-3
VS
V-5
V-BR-1
V-4
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo HPCDAE/ Pré-tratamento de biomassas com
explosão com dióxido de carbono
V-7
– Válvula de Bloqueio; V-BR-1, V-BR-2 – Válvulas Banho de Refrigeração (Controle da Circulação do Fluido
Refrigerante); VP-1 – Vaso de Pressão pequeno (6,57 mL); BL – Bomba para Líquidos (em condições atmosféricas); RT –
Rotâmetro; TO – Totalizador.
38
39
4. Formação de partículas via expansão rápida de solução supercrítica (Rapid
Extraction of Supercritical Solution – RESS) – Parâmetros importantes no
Processo
-
Pressão;
-
Temperatura;
-
Tempo de solubilização no fluido supercrítico (mais comum é o CO2);
-
Diâmetro do bico expansor (diâmetro interno do capilar);
-
Concentração do co-solvente - Relação entre co-solvente/solvente (cosolventes mais aconselháveis são: etanol e isopropanol);
-
Concentração do material de encapsulação - Relação entre material de
recheio/material de encapsulação.
A variação na Pressão e Temperatura da câmara de pré-expansão está
diretamente relacionada a variação na densidade do fluido supercrítico. Sendo a
seletividade do processo de formação de partículas ajustada através desta
mudança. Já a variação no diâmetro do bico expansor está diretamente associado
ao tamanhos das partículas formadas.
Segundo a experiência do LASEFI no processo de formação de partículas
encapsuladas via RESS, a temperatura exerce maior influência do que a pressão,
pois quando em contato com o CO2 supercrítico o material de encapsulação
40
absorve uma alta concentração do fluido o inchando e o fundindo, através da
redução da temperatura de fusão (em 10-15ºC). Uma vez fundido o material de
encapsulação, um maior aglomeramento das partículas é propiciado. Portanto, o
emprego de menores temperaturas é requerido para se obter partículas não
aglomeradas (Até 40 ºC para PEG 10.000, cujo ponto de fusão é 60 ºC).
Para processos em que o material de encapsulação possuem limitada solubilidade
no fluido supercrítico a adição de um co-solvente é capaz de aumentar a
solubilidade deste no fluido, produzindo, então, uma solução supercrítica do
material de recheio + material de encapsulação + fluido supercrítico + co-solvente.
Este processo é conhecido como RESS-N (Rapid Expansion of Supercritical
Solution with a Nonsolvent). O solvente orgânico, que na maioria das vezes é um
álcool, deve, além de aumentar a polaridade do fluido supercrítico, ser um não
solvente para as partículas formadas [No caso de utilizar PEG 10.000 como
material de encapsulação, etanol (em concentrações maiores que 25 % m/m) é
um bom não solvente para as partículas de PEG e um excelente co-solvente para
CO2 supercrítico, por exemplo].
Para processos de formação de partículas não encapsuladas a condição
necessária que as partículas devem obedecer é ser altamente solúveis no fluido
supercrítico. Caso tal condição não seja obedecida o processo conhecido como
41
SAS (Supercritical fluid Anti-Solvent – SAS) deve ser utilizado. Os processos de
formação de partículas não encapsuladas também são conhecidos como
processos de micronização, pois as partículas têm seu tamanho reduzido após a
rápida despressurização.
42
4.1 RESS (Procedimento)
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Se o objetivo é: i) formar partículas não encapsuladas de menor tamanho,
inserir o material a ser micronizado na câmara de pré-expansão (VP-1); ii) formar
partículas encapsuladas, inserir o material a ser encapsulado juntamente com um
material de encapsulação (biopolímero, ciclodextrina, etc.) na câmara de préexpansão [como muitos materiais de encapsulação possuem limitada solubilidade
em CO2 supercrítico, um determinado volume de co-solvente deve ser adicionado
também (RESS-N)];
3) Posteriormente, deve-se inserir a câmara de pré-expansão (VP-1) no sistema
de aquecimento do VP-1 desligado;
4) Conectar a tubulação VP-1/Entrada entre a válvula V-5 e o topo do VP-1;
5) Conectar a tubulação VP-1/Saída entre o fundo do VP-1 e a válvula V-6;
6) Conectar a tubulação VM/2 no orifício central do vaso separador (VP-2);
7) Verificar se as válvulas V-1, V-2, V-5 e V-6 estão fechadas e se a VM está
aberta;
8) Abrir a válvula do cilindro de CO2 e a válvula V-1 lentamente, observando se o
ponteiro do manômetro M-1 sobe suavemente;
43
9) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
10) Quando a pressão do CO2 for atingida, abrir a válvula V-5 para pressurizar a
câmara de pré-expansão (VP-1);
11) Ligar o controlador do sistema de aquecimento do vaso de pressão 1 (CAVP1) e da válvula micrométrica (CAVM) e programá-los para operar à temperatura
requerida do processo e a uma temperatura que evite congelamento da linha de
saída, respectivamente (esta temperatura não precisa ser muito alta (< 70 ºC),
pois o tempo de despressurização é muito rápido (menos de 4 segundos);
12) Quando a pressão e temperatura do sistema estiverem estabilizadas,
cronometrar o tempo estipulado para a solubilização dos solutos e fechar a válvula
V-5 (Este tempo pode oscilar de 20 minutos a 3 horas);
13) Decorrido o tempo de solubilização, abrir a válvula V-6 rapidamente
despressurizando o vaso VP-1, formando consequentemente partículas na parede
da câmara de precipitação (VP-2) em condições atmosféricas (Pressão e
Temperatura);
14) Desligar o banho BR e os controladores de aquecimentos CAVP-1 e CAVM e
fechar as válvulas V-1 e VC;
15) Coletar, pesar e guardar as partículas formadas em freezer à temperatura
igual ou menor que zero para posterior análise.
44
FLUXOGRAMA-RESS/RESS-N
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
V-7
BG
V-BPR
V-AR
M-3
BR
V-3
VS
V-5
V-4
V-BR-1
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo RESS/RESS-N
V-7
– Válvula de Bloqueio; V-BR-1, V-BR-2 – Válvulas Banho de Refrigeração (Controle da Circulação do Fluido
Refrigerante); M-4 – Manômetro; BA – Banho de Aquecimento; TP – Termopar; RT – Rotâmetro; TO – Totalizador.
45
46
5. Formação de partículas utilizando fluido supercrítico como anti-solvente
(Supercritical fluid Anti-Solvent – SAS) – Parâmetros importantes no Processo
-
Pressão;
-
Temperatura;
-
Vazão do anti-solvente (mais comum é o CO2);
-
Vazão da solução contendo as partículas;
-
Diâmetro do capilar coaxial interno (onde a solução contendo as partículas
é alimentada);
-
Concentração do material de encapsulação - Relação entre material de
recheio/material de encapsulação.
A variação na Pressão e Temperatura da câmara de precipitação está diretamente
relacionada a variação na densidade do fluido supercrítico. Sendo a seletividade
do processo de formação de partículas ajustada através desta mudança.
Segundo a experiência do LASEFI no processo de formação de partículas
encapsuladas, os parâmetros: i) Vazão do anti-solvente (mais comum é o CO2); ii)
Vazão da solução contendo as partículas;
iii) Diâmetro do capilar coaxial
interno (onde a solução contendo as partículas é alimentada); iv) Concentração do
material de encapsulação - Relação entre material de recheio/material de
encapsulação, são de igual importância aos parâmetros pressão e temperatura.
47
Para os processos de formação de partículas encapsuladas tanto o material de
recheio, quanto o material de encapsulação, devem ser altamente solúveis em um
solvente orgânico, o qual deve possuir uma alta miscibilidade no fluido supercrítico
(No caso de utilizar PEG 10.000 como material de encapsulação e CO2 como
fluido supercrítico, acetato de etila e diclorometano podem ser utilizados).
Assim como ocorre nos processos de formação de partículas via RESS, os
processos que produzem partículas não encapsuladas via SAS também são
conhecidos como processos de micronização, pois as partículas tem seu tamanho
reduzido após a sua precipitação, através da eliminação do solvente orgânico pelo
fluido supercrítico (anti-solvente).
Um processo muito similar à micronização via SAS, vem sendo desenvolvido
atualmente, o Supercritical Anti-solvent Extraction (SAE). Assim como o processo
SAS, este processo se utiliza da alta solubilidade de alguns solventes orgânicos
em fluidos supercríticos, porém ao invés de partículas em sua forma pura, extratos
vegetais contendo várias classes de compostos são empregados. Outra diferença
é que diferentemente do processo SAS, informações sobre alterações na
morfologia não são relevantes, pois partículas contendo uma mistura de
compostos são precipitados e não somente uma única. Aliado ao fato do processo
SAE formar partículas não encapsuladas, este processo pode proporcionar um
48
fracionamento de substâncias através de variações nos parâmetros: Pressão,
Temperatura, Vazão do anti-solvente (CO2 continua sendo o mais comum) e
Vazão da solução contendo a mistura de compostos (extratos metanólicos e
etanólicos têm sido utilizados em sua forma bruta ou parcialmente purificada).
49
5.1 SAS (Procedimento)
Previamente:
Se o objetivo é: i) formar partículas não encapsuladas de menor tamanho,
preparar uma solução homogênea contendo o material a ser micronizado em um
solvente orgânico; ii) formar partículas encapsuladas, preparar uma solução
homogênea contendo o material a ser encapsulado juntamente com um material
de encapsulação (biopolímero, ciclodextrina, etc.) em um solvente orgânico.
1) Ligar o banho de resfriamento (BR) e programá-lo para operar a – 10 ºC;
2) Ligar o banho de aquecimento (BA) e programá-lo para operar na temperatura
desejada;
3) Conectar a tubulação T no orifício central da câmara de precipitação (VP-2) e
suas ramificações na válvula V-5 e na bomba BL;
4) Conectar a tubulação VP-2 entre o orifício lateral (esquerda) da câmara de
precipitação (VP-2) e a válvula V-6;
50
5) Elaborar e inserir na extremidade da tubulação VP-2 que está dentro da câmara
de precipitação um cartucho de papel de filtro (10 μm) para evitar uma perda por
arraste das partículas formadas na câmara de precipitação;
6) Conectar a tubulação VM/1 na tampa de um frasco coletor lacrado (50 ou 100
mL) de massa conhecida, imerso em um banho de gelo;
7) Conectar a tubulação RT também na tampa do frasco coletor;
8) Verificar se as válvulas V-1, V-2, V-5, V-6 e VM estão fechadas;
9) Abrir a válvula do cilindro de CO2 e a válvula V-1 lentamente, observando se o
ponteiro do manômetro M-1 sobe suavemente;
10) Realizar a pressurização do CO2, abrindo a válvula VC (Válvula Compressor) e
V-BPR (Válvula Back Pressure Regulator), controlando o aumento da pressão
(Medição através do manômetro M-2);
11) Lacrar a câmara de precipitação (VP-2), conectando o termopar e o
manômetro M-4 ao vaso VP-2;
12) Quando a pressão do CO2 for atingida, abrir a válvula V-5 para pressurizar a
câmara de precipitação (VP-2);
13) Ligar o controlador do sistema de aquecimento da válvula micrométrica
(CAVM) e programá-los para operar a uma temperatura que evite congelamento
da linha de saída (esta temperatura depende da vazão de CO2 utilizada, podendo
oscilar entre 70 a 120 ºC);
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14) Quando a pressão e temperatura do sistema estiverem estabilizadas, abrir as
válvulas V-6 e VM cuidadosamente até que a vazão desejada seja alcançada
(Medição através do rotâmetro);
15) Estabilizada a vazão desejada, ligar a bomba para líquidos (BL) e programá-la
para alimentar a câmara de precipitação com a solução previamente preparada na
vazão desejada, pelo capilar interno do sistema coaxial (CO2 supercrítico passa
externamente) (Dependendo da pressão utilizada no vaso VP-2, a vazão
programada deve ser um pouco maior que a vazão desejada – utilizando uma
proveta verificar se há correspondência entre o valor de vazão programado e o
efetivamente desenvolvido, corrigindo-o, quando necessário);
16) Após o tempo pré-estabelecido de formação de partículas, desligar a bomba
BL para interromper a alimentação da solução;
17) Por um determinado tempo (de 20 minutos a 1 hora), continuar alimentando a
câmara de expansão somente com CO2 supercrítico a mesma vazão para eliminar
o solvente residual nas partículas formadas;
18) Decorrido o tempo de eliminação do solvente orgânico residual, desligar o
banho BR, o banho BA e o controlador de aquecimento CAVM e fechar as
válvulas V-5, V-1 e VC;
19) Despressurizar o vaso VP-2 empregando a mesma vazão utilizada no
processo de formação de partículas e eliminação do solvente residual, para não
52
provocar uma perda, por arraste, das partículas formadas na câmara de
precipitação;
20) Deslacrar a câmara de precipitação (VP-2), coletar, pesar e guardar as
partículas formadas em freezer à temperatura igual ou menor que zero para
posterior análise.
OBS: Quando ao invés de uma substância pura se emprega uma mistura
complexa, como, por exemplo, extratos metanólicos e etanólicos, o processo
recebe o nome de SAE. Portanto, para o desenvolvimento do processo SAE,
deve-se seguir procedimento exatamente igual ao descrito.
53
FLUXOGRAMA-SAS/SAE
CAVM
TP
M-2
M-1
M-4
VC
TO
CAVP-1
RT
V-6
VM
V-2
V-1
FL
V-7
BG
V-BPR
V-AR
M-3
BR
V-3
VS
Solução
V-5
V-4
V-BR-1
CO2
CO2
VP-1
V-BR-2
VP-2
BL
BA
Componentes da unidade que não são utilizados durante o processo SAS/SAE
V-7
– Válvula de Bloqueio; V-BR-1, V-BR-2 – Válvulas Banho de Refrigeração (Controle da Circulação do Fluido
Refrigerante).
54
55
ANEXOS
Densidade do CO2 em condições encontradas no laboratório
ρCO2
(kg/m3)
20 (ºC)
21
22
23
24
25
26
27
28
29
30
925 (mbar) 1.6784 1.6726 1.6668 1.6611 1.6554 1.6498 1.6442 1.6386 1.6331 1.6276 1.6222
926
1.6802 1.6744 1.6686 1.6629 1.6572 1.6516 1.6460 1.6404 1.6349 1.6294 1.6239
927
1.6820 1.6762 1.6704 1.6647 1.6590 1.6534 1.6477 1.6422 1.6366 1.6311 1.6257
928
1.6838 1.6780 1.6722 1.6665 1.6608 1.6551 1.6495 1.6440 1.6384 1.6329 1.6274
929
1.6857 1.6798 1.6740 1.6683 1.6626 1.6569 1.6513 1.6457 1.6402 1.6347 1.6292
930
1.6875 1.6816 1.6759 1.6701 1.6644 1.6587 1.6531 1.6475 1.6420 1.6364 1.6310
931
1.6893 1.6835 1.6777 1.6719 1.6662 1.6605 1.6549 1.6493 1.6437 1.6382 1.6327
932
1.6911 1.6853 1.6795 1.6737 1.6680 1.6623 1.6567 1.6511 1.6455 1.6400 1.6345
933
1.6929 1.6871 1.6813 1.6755 1.6698 1.6641 1.6585 1.6528 1.6473 1.6417 1.6363
934
1.6948 1.6889 1.6831 1.6773 1.6716 1.6659 1.6602 1.6546 1.6491 1.6435 1.6380
935
1.6966 1.6907 1.6849 1.6791 1.6734 1.6677 1.6620 1.6564 1.6508 1.6453 1.6398
936
1.6984 1.6925 1.6867 1.6809 1.6752 1.6695 1.6638 1.6582 1.6526 1.6470 1.6415
937
1.7002 1.6944 1.6885 1.6827 1.6770 1.6713 1.6656 1.6600 1.6544 1.6488 1.6433
938
1.7021 1.6962 1.6903 1.6845 1.6788 1.6731 1.6674 1.6617 1.6561 1.6506 1.6451
939
1.7039 1.6980 1.6922 1.6863 1.6806 1.6749 1.6692 1.6635 1.6579 1.6524 1.6468
940
1.7057 1.6998 1.6940 1.6882 1.6824 1.6767 1.6710 1.6653 1.6597 1.6541 1.6486
941
1.7075 1.7016 1.6958 1.6900 1.6842 1.6784 1.6727 1.6671 1.6615 1.6559 1.6503
942
1.7094 1.7035 1.6976 1.6918 1.6860 1.6802 1.6745 1.6689 1.6632 1.6577 1.6521
943
1.7112 1.7053 1.6994 1.6936 1.6878 1.6820 1.6763 1.6706 1.6650 1.6594 1.6539
944
1.7130 1.7071 1.7012 1.6954 1.6896 1.6838 1.6781 1.6724 1.6668 1.6612 1.6556
945
1.7148 1.7089 1.7030 1.6972 1.6914 1.6856 1.6799 1.6742 1.6686 1.6630 1.6574
Fonte: NIST (http://webbook.nist.gov)
56
Densidade do CO2 em condições sub/supercríticas (Pc = 73,3 bar; Tc = 31ºC)
ρCO2
(kg/m3)
30 (ºC)
35
40
45
50
55
60
70 (bar)
266.56 220.08 198.02 183.20 172.01 163.03 155.53
80
701.72 419.09 277.90 241.05 219.18 203.64 191.62
90
744.31 662.13 485.50 337.51 285.00 255.55 235.39
100
771.50 712.81 628.61 498.25 384.33 325.07 289.95
110
792.10 743.95 683.52 603.15 502.64 414.90 357.79
120
808.93 767.07 717.76 657.74 584.71 504.51 434.43
130
823.25 785.70 743.04 693.65 636.12 571.33 505.35
140
835.79 801.41 763.27 720.47 672.17 618.45 561.37
150
846.98 815.06 780.23 741.97 699.75 653.50 604.09
160
857.12 827.17 794.90 759.98 722.09 681.12 637.50
170
866.41 838.09 807.87 775.53 740.88 703.82 664.59
180
875.00 848.04 819.51 789.24 757.12 723.08 687.25
190
883.00 857.21 830.09 801.53 771.45 739.81 706.68
200
890.50 865.72 839.81 812.69 784.29 754.61 723.68
210
897.56 873.67 848.81 822.91 795.94 767.88 738.78
220
904.23 881.15 857.20 832.36 806.61 779.93 752.38
230
910.57 888.20 865.07 841.16 816.46 790.97 764.73
240
916.61 894.88 872.48 849.39 825.62 801.17 776.07
250
922.38 901.23 879.49 857.14 834.19 810.65 786.55
260
927.91 907.29 886.14 864.46 842.25 819.52 796.30
270
933.22 913.08 892.48 871.40 849.85 827.85 805.42
280
938.32 918.64 898.53 878.00 857.05 835.70 813.98
290
943.24 923.97 904.33 884.30 863.90 843.15 822.06
300
947.98 929.11 909.89 890.33 870.43 850.22 829.71
350
969.56 952.29 934.81 917.12 899.23 881.17 862.94
400
988.31 972.26 956.07 939.75 923.32 906.77 890.14
Fonte: NIST (http://webbook.nist.gov)
57
Viscosidade da água em condições sub/supercríticas (Pc = 221.2 bar; Tc = 374ºC)
Viscosidade
água (cP)
30 (ºC)
40
50
60
70
80
90
100
120
150
200
250
300
350
400
1 (bar)
0.79735
0.65298 0.54685 0.46640 0.40389 0.35435 0.31441 0.01227 0.01302 0.01418 0.01617 0.01822 0.02029 0.02237 0.02445
20
0.79717
0.65313 0.54718 0.46682 0.40437 0.35485 0.31492 0.28225 0.23253 0.18285 0.13443 0.01785 0.02007 0.02225 0.02440
40
0.79700
0.65330 0.54753 0.46727 0.40487 0.35537 0.31545 0.28279 0.23305 0.18335 0.13493 0.10612 0.01988 0.02216 0.02437
50
0.79692
0.65339 0.54771 0.46750 0.40512 0.35564 0.31572 0.28305 0.23331 0.18360 0.13518 0.10640 0.01979 0.02212 0.02436
60
0.79685
0.65348 0.54789 0.46773 0.40537 0.35590 0.31599 0.28332 0.23357 0.18385 0.13542 0.10668 0.01973 0.02210 0.02437
70
0.79677
0.65357 0.54807 0.46796 0.40563 0.35616 0.31625 0.28359 0.23384 0.18411 0.13567 0.10696 0.01967 0.02209 0.02438
80
0.79671
0.65367 0.54825 0.46818 0.40588 0.35643 0.31652 0.28386 0.23410 0.18436 0.13592 0.10723 0.01965 0.02209 0.02441
90
0.79664
0.65377 0.54844 0.46842 0.40613 0.35669 0.31679 0.28412 0.23436 0.18461 0.13616 0.10751 0.08606 0.02211 0.02444
100
0.79658
0.65387 0.54863 0.46865 0.40639 0.35696 0.31706 0.28439 0.23462 0.18486 0.13641 0.10778 0.08646 0.02215 0.02448
120
0.79648
0.65408 0.54900 0.46911 0.40690 0.35749 0.31760 0.28493 0.23514 0.18536 0.13689 0.10831 0.08723 0.02230 0.02461
150
0.79635
0.65441 0.54958 0.46982 0.40767 0.35829 0.31840 0.28573 0.23592 0.18610 0.13761 0.10910 0.08833 0.02293 0.02493
200
0.79620
0.65500 0.55056 0.47101 0.40896 0.35962 0.31975 0.28707 0.23722 0.18733 0.13880 0.11039 0.09005 0.06930 0.02603
250
0.79615
0.65565 0.55158 0.47222 0.41026 0.36096 0.32109 0.28840 0.23851 0.18856 0.13998 0.11164 0.09164 0.07276 0.02917
300
0.79619
0.65636 0.55264 0.47345 0.41158 0.36231 0.32244 0.28974 0.23980 0.18977 0.14113 0.11285 0.09315 0.07545 0.04393
350
0.79633
0.65712 0.55372 0.47470 0.41291 0.36366 0.32379 0.29107 0.24109 0.19098 0.14227 0.11403 0.09457 0.07773 0.05579
400
0.79655
0.65793 0.55484 0.47597 0.41425 0.36502 0.32515 0.29241 0.24237 0.19218 0.14340 0.11519 0.09593 0.07974 0.05937
Fonte: NIST (http://webbook.nist.gov)
58
Valores da constante dielétrica de alguns solventes orgânicos
P = 1 bar 20 (ºC)
P = 1 bar 20 (ºC)
Água
80.20
25
40
Água
80.20
Metanol
33.64
Etanol
25.16
Acetona
21.13
Butanol
18.19
Hexano
1.8
60
80
100
78.50 73.12 66.62 60.58 55.10
150 200 250 300 350 400
40
35
28
20
18
18
*Valores aproximados (retirados de figuras)
Fonte: Sites da internet, artigos e livros.
Entre 150 e 300 ºC, em altas pressões, a água é capaz de dissolver muitos
compostos polares, pois sua constante dielétrica é similar a dos solventes Metanol
e Etanol, em contrapartida acima de 300 oC, a água pressurizada é capaz de
dissolver muitos compostos apolares. Mais diferente ainda é a água quando a
pressão for igual ou maior de 221.2 bar e a temperatura maior do que 374 ºC
(pressão e temperatura críticas): a água se torna um fluído supercrítico, tendo sua
constante dielétrica oscilando entre 30 e 2. Nestas condições, a água reúne
propriedades de gás (tal como a densidade) e de líquido (capacidade de
dissolução). Além de dissolver substâncias polares e iônicas, a água supercrítica é
capaz de dissolver praticamente todos os compostos apolares, pois a constante
dielétrica é similar a do hexano (1.8). Adicionalmente, nestas condições a água
supercrítica é um excelente meio reacional, podendo ser utilizada, por exemplo, na
destruição de lixos tóxicos, hidrólise de biomassa, etc.
59