Holistic valorization of rapeseed meal utilizing green solvents extraction and

biopolymer production with Pseudomonas putida

Phavit Wongsirichota, Maria Gonzalez-Miquela,b and James Winterburna*

a School of Chemical Engineering and Analytical Science, The University of Manchester,

Oxford Rd, Manchester, M13 9PL, United Kingdom

b Departamento de Ingenieria Quimica Industrial y del Medio Ambiente, ETS Ingenieros

Industriales, Universidad Politécnica de Madrid, Calle de José Gutiérrez Abascal 2,

Madrid, 28006, Spain

* Corresponding author

Email: James.Winterburn@manchester.ac.uk, Phone: +44 (0) 161 306 4891

Abstract:

A rapeseed meal (RSM) valorization scheme was developed to utilize valuable fractions

including phenolics (mainly sinapic acid), proteins and polysaccharides. This involved

solvent extraction of phenolics, alkali extraction of proteins and fermentation of residual

polysaccharide using Pseudomonas putida. For the first time, sustainable Deep eutectic

solvents (DESs) were used in the extraction of rapeseed meal phenolics. With yields up

to 85.69 % wt. DESs were able to outperform methanol (59.54 % wt.) at sinapic acid

extraction. As shown by Conductor like Screening Model for Real Solvents (COSMO-

RS), this is because DESs have greater capacity for H-bonding.

Post-extraction RSM hydrolysate was shown to be a viable media for cultivation of P.

putida with maximum cell dry weight of 4.89 g l−1 for DES-extracted RSM. 8-carbon and

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10-carbon chain length polyhydroxyalkanoate biopolymers were synthesized.

Biopolymer accumulation was reduced in RSM derived media due to high nitrogen

concentration. These findings are beneficial for the development of a sustainable

biorefining scheme based on rapeseed meal.

Highlights:

• Valorization scheme investigated for rapeseed phenolics, proteins and

polysaccharides

• Deep eutectic solvents outperformed methanol in extracting rapeseed meal

phenolics

• Rapeseed meal hydrolysate shown to be viable media for Pseudomonas putida

• Successful synthesis of medium chain-length polyhydroxyalkanoates (8- and 10-

carbon)

Keywords: Rapeseed meal, valorization; phenolics; deep eutectic solvents;

polyhydroxyalkanoates

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1. Introduction

Over 70 million tons of rapeseed is grown annually for oil production (USDA, 2018),

by mass, over 60% of the grain will end up as waste called rapeseed meal (RSM)

(D'Avino et al., 2015). While, RSM can be sold as animal feed, this is limited by anti-

nutritional compounds such as glucosinolates. In order to reduce these compounds,

further processing, such as methanol extraction will be needed (Qian et al., 2013). More

importantly, RSM contains protein, phenolics, and polysaccharides, which could be

potentially valorized.

RSM phenolics are of major interest due to their significant anti-oxidative properties

(Nowak et al., 1992). Consequently, derivatives of RSM phenolics, such as 4-

vinylsyringol could potentially be used in the food industry due to their anti-oxidative

capabilities and flavors (Harbaum-Piayda et al., 2010). RSM phenolics’ anti-

inflammatory, anti-mutagenic properties and ability to improve drug permeability could

also lead to medical applications (Vuorela et al., 2005).

The major phenolic compounds in RSM are phenolic acids in both free and esterified

forms. Sinapic acid makes up the majority of the phenolics, mainly existing as the

choline ester, sinapine. Other phenolic acids include trans-ferulic, p-hydroxybenzoic,

coumaric and syringic acids (Naczk et al., 1998).

High sinapic acid yields have been achieved by extractions utilizing traditional

solvents such as methanol and acetone (Das Purkayastha et al., 2013). However, the

use of these conventional solvents has inherent disadvantages due to their toxic,

flammable, and/or corrosive properties, which can negatively impact operator safety and

the environment (Anastas and Eghbali, 2010).

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There is growing interest in the use of deep eutectic solvents (DESs) for extraction of

bioactive compounds from various biomass such as sea buckthorn leaves (Cui et al.,

2018), green tea (Jeong et al., 2017), cannabis (Křížek et al., 2018) and citrus by-

products (Ozturk et al., 2018a; Ozturk et al., 2018b) DESs are mixtures comprising of a

hydrogen bond acceptor (HBA) and a hydrogen bond donor (HBD). HBAs are typically a

quaternary ammonia salt. HBD can range widely from sugars to alcohols and organic

acids. However, despite their potential, DESs have not yet been applied to the

extraction of RSM phenolics.

Aside from the phenolics, RSM also contains potentially valuable proteins and

polysaccharides. Whilst there exists an extensive literature on RSM proteins, there is

less data on the utilization of the polysaccharides. RSM polysaccharides have been

utilized in both solid-state fermentation (Ebune et al., 1995) and fermentation with

hydrolyzed media (Chen et al., 2011). To the best of the authors’ knowledge, RSM-

derived polysaccharides have not been used in biopolymer production, warranting

further investigation.

An ideal organism for fermenting RSM should be able to metabolize the various

saccharides present in cellulose and hemicellulose. A good candidate is Pseudomonas

putida, which can metabolize a wide range of carbon sources, including saccharides

(Stanier et al., 1966). These saccharides can then be converted via de novo fatty acid

synthesis to polyhydroxyalkanoates (PHA) (Prieto et al., 2016). PHAs are polyesters

produced for energy storage whose accumulation is typically triggered by a limitation of

nutrients such as nitrogen in the presence of excess carbon substrate (Kachrimanidou

et al., 2014). Due to their biodegradability and thermoplastic properties, PHAs have a

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wide range of potential applications from substitute for conventional plastics to

biomaterials (Singh et al., 2015). Waste biomass have often been used as a carbon

source for PHA production, such as palm oil effluent (Mumtaz et al., 2010) and waste

plant oils (Ciesielski et al., 2015).

Currently, integrated RSM valorization of all three components remain largely

unexplored, with the only example being Li and Guo (2017). As fermentation was not

conducted, the viability of sugars from post-extraction RSM remains to be proven.

In this study, extractions of sinapic acids were conducted using a range of DESs.

Conductor like Screening Model for Real Solvents (COSMO-RS) simulations were used

to better understand the underlying molecular interactions. Finally, bioreactor cultures of

P. putida were grown using post-extraction RSM as media. This study presents the first

proof-of-concept study of the full valorization of RSM components, incorporating

extraction of high-value components using sustainable DESs, as well as biopolymer

production from the residual saccharides.

2. Materials and Methods

2.1 Materials

Defatted RSM was supplied by Cargill PLC (UK), which was passed through a 1 mm

sieve (Endecotts, UK) prior to use. P. putida KT2440 was procured from DMSZ,

Germany and stock cultures were prepared according to the supplier’s instructions.

Chemical used were procured from Sigma-Aldrich (UK), Fisher Scientific (UK), Alfa

Aesar (UK) VWR International (UK) and Merck (UK), which were as follows: Nutrient

agar II, sinapic acid (≥98 %), glucose (≥99 %), galactose (≥99 %), arabinose (≥98 %),

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xylose (≥99 %), 3-hydroxydecanoic acid (≥98 %), 3-hydroxyoctanoic acid (≥97 %),

octanoic acid (≥99 %), decanoic acid (≥98 %), dodecanoic acid (≥98 %), Folin-ciocalteu

reagent (2 N), sodium carbonate (≥99 %), ethylene glycol (≥99.8 %), trisodium citrate

(≥99 %), K2HPO4 (≥98 %), KH2PO4 (≥98 %), Na2HPO4 (≥98.5 %), FeSO4·7H2O (≥99 %),

CaCl2·2H2O (≥99 %), MnCl2·4H2O(≥98 %), CoSO4·7H2O (≥99 %), CuCl2·2H2O (≥99 %),

MgSO4·7H2O (≥98 %), methanol (≥99.8 %), NaNO3 (≥99 %), , NaOH (98 %), NH4Cl (≥98

%). H2SO4 (98 %), HCl (1 M), glacial acetic acid, choline chloride (99 %), glycerol (99 %)

and NaCl (99 %).

2.2 RSM phenolics characterization

Consecutive extractions were conducted using methanol (3 extractions) followed by

acetone (final extraction). All extractions were performed in a water bath for 2 hours at

40 ºC and stirred at 1500 rpm. The solid-to-liquid ratio was 1 g per 10 ml. Samples were

centrifuged at 3000 rpm for 20 minutes using a Sigma 6-16S (SciQuip, UK). The

supernatants were then collected and stored at −20 ºC prior to further analysis.

Colorimetric measurement of total phenolics was conducted using the Folin-ciocalteu

method based on Szydłowska-Czerniak et al. (2011).

The supernatant was hydrolyzed using a method based on Naczk et al. (1992).

Measurements of individual phenolic acids were conducted using high performance

liquid chromatography (HPLC) using an UltiMate® 3000 system (Thermo Fisher

Scientific, UK) and a 150 mm Nucleosil® C18 column (Macherey-nagel, Germany).

HPLC method used was based on Cai and Arntfield (2001). Sample injection volume of

5 µl was used, and sinapic acid was detected using UV absorbance at 330 nm. Other

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phenolic acids, including syringic acid, p-coumaric acid, ferulic acid and benzoic acid

were measured at 250 and 278 nm.

2.3 DESs preparation

DESs were prepared at ratios shown in table 1. The components were mixed for 24

hours between 250 - 500 rpm and 60 ºC. The DESs were then dried at 50 ºC in a

VT6025 vacuum oven (Thermo Fisher Scientific, UK) for 24 hours. Aqueous DESs were

prepared by addition of distilled water, at a DES to water ratio of 7:3 v/v (Nam et al.,

2015).

Hydrogen Bond Acceptor Hydrogen Bond Donor Molar ratio Abbreviations

Choline chloride Glucose 1 : 1 ChCl: GC (1: 1)

Choline chloride Glucose 1.5 : 1 ChCl: GC (1.5: 1)

Choline chloride Glucose 2 : 1 ChCl: GC (2: 1)

Choline chloride Glycerol 1 : 2 ChCl: Gly (1: 2)

Choline chloride Glycerol 1 : 1.5 ChCl: Gly (1: 1.5)

Choline chloride Glycerol 1 : 1 ChCl: Gly (1: 1)

Choline chloride Ethylene Glycol 1 : 3 ChCl: EG (1: 3)

Choline chloride Ethylene Glycol 1 : 2 ChCl: EG (1: 2)

Choline chloride Ethylene Glycol 1 : 1.5 ChCl: EG (1: 1.5)

Table 1 Composition of DESs used

2.4 Sinapic acid solubility

Sinapic acid was added in excess to 2 ml solvents. The samples were stirred for 2

hours at 1000 rpm and 30 ºC. Analysis of sinapic acid concentration was conducted as

described in the previous section.

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2.5 Solvent extraction of phenolics from RSM

Extractions of RSM phenolics were conducted with aqueous DES and aqueous

methanol, at 40 ºC, 50 ºC and 60 ºC, for 2 hours at 1000 rpm. The solid-to-liquid ratio

was 1 g per 10 ml. After extraction, the samples were centrifuged at 3000 rpm for 20

minutes and the supernatant removed. HPLC analysis was conducted as described

above. Sinapic acid yields with respect to total sinapic acid were calculated using

equation (1).

Sinapicacid yield (%wt )= Sinapicacid∈sample (g l−1 )×100 %Sinapic acid∈RSM (gg−1 )×Solid loading (g l−1)

(1)

Scaled-up extraction of RSM using methanol and the best performing DES (ChCl: Gly,

1:1) was conducted using 250 g RSM in 2.5 l at 60 ºC and 200 rpm. The residues were

separated using a Pyrex coarse grain Buchner funnel (Corning, USA). The DES

extracted RSM were then washed three times using 1 l of distilled water at 180 rpm for

5 minutes. The post-extraction RSMs were vacuum dried for 48 hours at 50 ºC.

2.6 Measurement of DES viscosities

Viscosity measurements were conducted using an DMA™ 4500 M density meter

equipped with an AMVn Automated Micro Viscometer (Anton Paar, UK).

2.7 Preparation of fermentation media

Synthetic media formulation was based on previous works conducted with P. putida

by Hartmann et al., (2006); Le Meur et al., (2012) and Davis et al., (2013). Preculture

media was prepared with 2.9 g l−1 trisodium citrate as a carbon source. Additional

nutrients were as follows: 7.5 g l−1 K2HPO4, 3.7 g l−1 KH2PO4, 2.38 g l−1 Na2HPO4, and

0.896 g l−1 NH4Cl. Sterilization was done via autoclaving at 121 ºC for 20 minutes (BMM

Weston, UK). After autoclaving, filter sterilized solutions of other trace elements were

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added at a concentration of 1 ml l−1. The trace element solution consisted of

FeSO4·7H2O (2.78 g l−1), CaCl2·2H2O (1.47 g l−1), MnCl2·4H2O (1.98 g l−1), CoSO4·7H2O

(2.81 g l−1), CuCl2·2H2O (0.17 g l−1) in 1 M HCl. 1 ml l−1 of filter sterilized MgSO4·7H2O

solution (1 M) was also added.

For the control experiment, a similar media was prepared which utilized glucose,

xylose, galactose and arabinose as a carbon sources, at concentrations of 5.5, 1.5, 3.5,

and 5 g l−1, respectively. NH4Cl (0.896 g l−1) was used as a nitrogen source, resulting in

an initial carbon to nitrogen ratio of 26 g g−1. The sugar content was selected based on

results from 100 ml scale diluted hydrolysis of RSM using the procedure in the following

section.

2.8 RSM hydrolysis

RSM hydrolysis was needed to produce monosaccharides for use as fermentation

carbon sources. Prior to hydrolysis, proteins were removed from post-phenolic

extraction RSM by using 0.4% w/v sodium hydroxide, as per a method by Klockeman et

al. (1997). Subsequently, the RSM was washed with distilled water and centrifuged. The

RSMs were vacuum dried for 48 hours at 50 ºC. Diluted acid hydrolysis was conducted

on 200 g of RSMs using 2 l of 6 % wt. H2SO4. Hydrolysis was conducted for 1 hour at

121 ºC within an autoclave. Hydrolysate was neutralized to pH 7 ± 0.5 using solid

NaOH. Phosphates and trace elements were added at the same concentration as the

control. The hydrolysates were filter sterilized using a 0.45 µm Nalgene™ Rapid-Flow™

filter (Fisher Scientific, UK).

2.9 Fermentation of P. putida

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Cells were initially grown on nutrient agar II, for 24 hours at 30 ºC. Pre-cultures of 200

ml were grown at 30 ºC and 200 rpm for 24 hours. The bioreactor working volume and

total volume were 2 l and 3 l, respectively (Applikon, UK). Temperature was maintained

at 30 ± 2 ºC. pH was maintained at 7 ± 1 using 3 M HCl and 3 M NaOH solutions.

Struktol J647 was used as antifoam. Dissolved oxygen was maintained above 30%

during fermentation using cascade control with stirrer speed between 600 to 1500 rpm.

2.10 Measurement of bacterial growth

Bioreactor samples were centrifuged at 13,400 rpm for 5 minutes (Eppendorf,

Germany). The supernatant was removed for further analysis. The cell pellet was

resuspended in 0.7 % wt. NaCl solution. Optical density at 600 nm (OD600) was

determined using a UV mini 1240 UV spectrophotometer (Shimazdu, UK). For cell dry

weight measurements, centrifuged cell pellets were washed with distilled water and

dried at 70 ºC.

2.11 HPLC analysis of sugars

HPLC used was an UltiMate® 3000 system with a RefractoMax 520 refractive index

detector (Thermo Fisher Scientific, UK). The columns used were an Aminex HPX-87P

and Aminex HPX-87H (Bio-rad, UK) with an isocratic elution using 0.6 ml min−1 of water

and 0.05 mM H2SO4, respectively. Sample injection volume was 10 µl and column

temperature was 50 ºC.

2.12 Total nitrogen analysis

A TOC-VCPH Total Organic Carbon analyzer with a TNM-1 TN analyzer unit

(Shimadzu, UK) was used for total nitrogen analysis, standard solutions of NaNO 3 was

used for calibration.

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2.13 GC-FID of PHAs

Methanolysis was used to treat the dried cells based on Lageveen et al. (1988). The

derivatized PHA was detected using Gas Chromatography with a Flame Ionization

Detector (GC-FID). GC-FID was conducted on an Ultimate 1300 GC system (Thermo

Fisher Scientific, UK), using a Zebron ZB-SemiVolatiles Capillary GC Column, 30m x

0.25mm x 0.25µm (Phenomenex, UK). Sample injection volume of 1 µl was used.

Helium was employed as a carrier gas at 1 ml min−1 with a split ratio of 50:1.

The following oven temperature profile was used: Initially, the oven was maintained at

100 ºC for 3 minutes, followed by heating at a linear gradient of 25 ºC min−1 until 320 ºC,

and finally the oven was held at 320 ºC for 2 minutes. Inlet and detector temperatures

were at 300 ºC. Concentrations of different chain-length PHAs were determined by

comparison to standard solutions of methanolized 3-hydroxydecanoic and 3-

hydroxyoctanoic acids.

2.14 Fermentation yield coefficients

Yield coefficients were calculated using equations (2) to (4). As the amino acids can

also be catabolized by P. putida (Radkov and Moe, 2013), equation 3 which took the

nitrogen source into account was also used.

Y XC=CDW t−PHAt

C (2)

Y XC+N

=CDW t−PHA t

C+N (3)

Y PHAGC

=PHA t

GC (4)

Where:

X is the residual cell weight at time t (g l−1)

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C is the carbon source consumed by time t (g l−1)

CDW t is the cell dry weight at time t (g l−1)

PHA t is the PHA content in the culture at time t (g l−1)

N is the nitrogen source consumed by time t (g l−1)

GC is the glucose consumed by time t (g l−1)

For RSM cultures, the amount of nitrogen source consumed (i.e. proteins) was

estimated using equation (5).

For RSM :N=TN t−TN 0

NRSM(5)

Where:

TN t is the total nitrogen at time t (g l−1)

TN 0 is the total nitrogen at 0 h (g l−1)

N RSM is the mass ratio of nitrogen in RSM proteins

A value of 0.1384 was used for N RSM, based on the prominent amino acids in RSM

reported by Tzeng et al.(1988).

The specific growth rate, µ, was calculated based on equation (6) (Blanch and Clark,

1996).

lnCDW t

CDW 0=µ(t−t lag) (6)

Where:

CDW 0 is the cell dry weight at 0 h (g l−1)

t lag is the duration of the lag phase (h)

2.15 COSMO-RS

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Conductor like Screening Model for Real Solvents (COSMO-RS) was utilized to

calculate thermodynamic properties of sinapic acid in solution. Geometric optimization

and COSMO file calculation for sinapic acid was performed using Gaussian 09 at the

BVP86/TZVP/DGA1 quantum chemical level (Frisch et al., 2016). COSMO files for other

molecules were taken from COSMO-RS database. COSMOtherm version C3.0 release

17.05 was used at the corresponding parametrization (BP_TZVP_C30_1701) to

calculate σ-profiles, excess enthalpies and energetic contributions from hydrogen

bonding, misfits/electrostatic interactions and van der Waals forces (Klamt and Eckert,

2000). DESs were treated separately as choline cation, chloride anion, and the H-bond

donor, following the electroneutral approach (Hizaddin et al., 2014).

3. Results and discussion

3.1 Phenolic extractions

3.1.1 Choice of DES components

The main criteria when assessing the viability of the solvents for an integrated RSM

valorization process were solvent performance and the possible impact of residual

solvent. Importantly, toxicity needed to be considered due to potential impact on the

subsequent fermentation. Fortunately, there are many potential combinations of HBD

and HBA that can produce DES, many of which could be benign to the fermentation

process. Typically, HBAs are quaternary ammonium salts such as choline chloride

(Duan et al., 2016). In fact, choline compounds can actually be metabolized by

Pseudomonas species, including P. putida.(Galvão et al., 2006) Therefore, the use of

choline chloride based DESs was deemed appropriate for phenolic extraction.

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The choice of HBDs can vary widely from organic acids, polyols and sugars (Dai et

al., 2013). Glycerol (Abbott et al., 2007; Ozturk et al., 2018a) and glucose (Hayyan et

al., 2013) were chosen due to their prevalence within the literature for polyols and

sugar-based DESs, respectively. They also have additional benefits stemming from

their renewable nature and the ability of P. putida strains for both glucose and glycerol

catabolism (Stanier et al., 1966). Ethylene glycol (EG) was chosen to compare

performance to the chosen DESs, due to high extraction performance demonstrated

within literature (Abbott et al., 2007; Ozturk et al., 2018b). Finally, methanol (MeOH)

was used for comparison, as it is a commonly utilized traditional solvent for extraction of

RSM phenolics (Das Purkayastha et al., 2013).

3.1.2 RSM phenolic content

During the characterization process, sinapic acid was confirmed to be the most

abundant phenolic compound within the RSM. Total sinapic acid in the RSM was 8.97

mg g−1. This was much higher than the other phenolic acids such as syringic acid, p-

coumaric acid, ferulic acid and benzoic acid, which were not detected in significant

amounts. Based on the Folin–Ciocalteu method, the total phenolics in RSM were found

to be 16.14 mg g−1 (sinapic acid equivalent). At approximately 60 % wt., the proportion

of sinapic acid within total phenolics was found to be lower than RSM within literature,

which ranges from 70 to 97 % (Naczk et al., 1998; Naczk et al., 1992). However, the

amount of sinapic acid agrees with literature values, which can range significantly from

approximately 5.55 mg g−1 to 11.80 mg g−1 (Khattab et al., 2010; Vuorela et al., 2003).

Variations are likely due to different sources of RSM both in terms of agricultural source

and rapeseed processing method.

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3.1.3 Preliminary screening of DESs for phenolic extraction

A preliminary screening was initially conducted on DES at 40 ºC (Fig. 1a). With yields

ranging between 67.5 to 72.9 % wt., extractions using ChCl: Gly and ChCl: EG DESs

demonstrated slightly better sinapic acid yield than methanol at 59.5 % wt. The

difference became more pronounced once the extraction temperature was raised, with

sinapic acid yields between 78.9 and 85.7 % wt. for DES ChCl: Gly and ChCl: EG at 60

ºC, while yields for methanol remained constant with temperature. The correlation

between extraction temperature and yields for most of the DESs used was likely due to

improved mass transfer as a result of decreasing viscosity with increasing temperature.

To avoid degradation of the phenolics the temperature was not increased beyond 60 ºC.

Figure 1: a) Sinapic acid yields from RSM using aqueous solvents, b) Solubility of

sinapic acid Error bars indicate standard error from triplicates

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The choice of HBD played an important role in the extraction efficiency. The maximum

phenolic yields for ChCl: GC (1: 1) was approximately 48% wt. at 60°C. At the same

temperature, ChCl: Gly (1: 1) was above to achieve at yield of 85 % wt. This agrees with

similar findings from Nam et al. (2015) regarding flavonoids extraction, where ChCl: Gly

also out performed ChCl: GC. This was due to the effect of HBD choice on both the

polarity and viscosity of the resulting DES. Glucose also contains a higher number of

hydroxy groups compared to glycerol and ethylene glycol. This means there is higher

probability of solvent-solvent interaction in ChCl: GC which could also reduce the

solvent-solute interaction, resulting in lower sinapic acid yields. Interestingly, changing

the proportion of HBD to HBA did not have significant impact on the sinapic acid yield

for the DESs. Further comparison with literature values for RSM is made difficult

because the majority of the papers utilized Folin–Ciocalteu data as the main

measurement, or because mg g−1 RSM values were used, which will be impacted by the

native amount present in different sources of RSM.

The solubility of pure sinapic acid was also assessed to determine if there are

differences in performance compared to the biomass (Fig. 1b). For pure sinapic acid

with both ChCl: Gly and ChCl: GC, solubility decreased with increasing proportion of

HBD. For example, sinapic acid concentrations were 1.56 and 0.65 g l−1 for ChCl: Gly (1:

1) and (1: 2) respectively. This was not the case for the EG based DESs. This was

mainly due to an increase in viscosity as the proportion of HBD increases. At 16.81 g l−1,

the solubility of sinapic acid in methanol was in fact far higher than the DES which

ranged between 0.27 to 1.66 g l−1. The much poorer performance of methanol on RSM

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could be because other compounds with RSM are being preferentially dissolved or co-

extracted, reducing extraction selectivity with respect to phenolics.

COSMO-RS simulations were conducted to better understand the interaction between

solute and solvents and potentially explain the trends observed in Fig. 1. In COSMO-

RS, the type of intermolecular behavior can be inferred from the molecules σ-profile

(Klamt and Eckert, 2000). The σ-profiles of a molecule can be divided into three

regions, displaying H-bond donor (σ < −0.0082 e Å−2), non-polar (−0.0082 e Å−2 < σ <

0.0082 e Å−2), and H-bond acceptor (0.0082 e Å−2 < σ) behaviours (González et al.,

2018). Comparison of the solute and solvent σ-profile can determine the potential of

solubility. If both histograms are complementary, for example, if one has a peak

indicating H-bond donor behavior, while the other has a peak indicating H-bond

acceptor behavior, there is a potential for good solubility (Lapkin et al., 2010). While this

analysis of σ-profile is typically conducted on liquid-liquid systems, such treatment of

solute-solution systems has precedence within literature such as those by Lapkin et al.

(2010), Gonzalez et al. (2018) and Pereira et al. (2016).

σ-profile of sinapic acid with respect to the solvent’s components are shown in Fig. 2a

and 2b. Sinapic acid displays significant peaks in the H-bond donor and H-bond

acceptor regions. The H-bond acceptor peak extended to σ of approximately 0.017 e

Å−2, displaying a maximum at σ of 0.011 e Å−2. This is due to the contributions from the

carboxylic acid and phenolic functional groups, as well as the oxygen atoms of the

hydroxyl groups. This is also demonstrated by the visualization of the σ surface shown

in Fig. 2d.

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Meanwhile, the H-bond donor peak extended to σ of approximately −0.022 e Å −2.

While maximum value of p(σ) for the H-bond acceptor peak is lower at around 2

compared to 4.5 for the H-bond donor peak, the broadness of the charge distribution

means that the H-bond donor behavior is no less significant. Again, both the phenolic

and carboxylic acid functional groups are the cause of this behavior, as shown in Fig. 2.

The potential for sinapic acid to act as a H-bond donor or acceptor has been seen in

literature. For example, Sinha et al. (2015) reported a range of H-bonding behaviors

between sinapic acid and a number of APIs. This was caused by either the carboxylic

acid acting as both H-bond donor and acceptor, or by the phenolic functional group

acting as a H-bond donor (Sinha et al., 2015).

The solubility of sinapic acid could be directly impacted by the ease of formation of H-

bonds between the solute and the solvents. This had been previously demonstrated by

orange peels phenolics, where increased H-bonding improved the solubility of orange

peel phenolics in DES, although this can be hindered by kinetic effects (Ozturk et al.,

2018b). Fig. 2 shows that as expected the DESs components displays significant H-

bond donor and acceptor behaviors. Among the designated HBDs, glucose is the most

polar as it has comparatively more polar functional groups (including hydroxymethyl and

hydroxyl groups) relative to glycerol and ethylene glycol which have similar σ-profiles.

Additionally, the polarity from the choline cation or the chloride anion may also aid

sinapic acid solubility. The σ-profile of methanol contains much smaller peaks in both H-

bond donor or acceptor regions compared to the DES components.

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Figure 2: σ-profile of sinapic acid with respect to solvents components a) Methanol and

ChCl, b) DES H-bond donors, c) molecular structure of sinapic acid, white: H-atom, red:

O-atom, green, d) visualization of σ surfaces for sinapic acid, red and blue denote more

extreme σ values, red: positive, blue: negative, green: neutral

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In addition to the σ-profiles, excess enthalpy (ΔH) was calculated to directly compare

the molecular interaction within each solvent system. These calculations considered the

molar proportion of each component, rather than just an individual atom as with σ-

profiles. Additionally, it also took into account multiple types of molecular interactions:

van der Waals forces, electrostatic misfits and H-bonding (Klamt and Eckert, 2000).

High solubility of a solute is indicated by high exothermic behavior of the system, i.e.

more negative values of ΔH (Gonzalez-Miquel et al., 2013). This is shown in Fig. 3,

which in all cases, the maximum ΔH of the sinapic acid - solvent system is a net

negative for both DESs and methanol, indicating favorable intermolecular interactions

that enhance solubility. Maximum ΔH for DES ranges from −1.00 to −2.02 kcal mol−1,

much greater than the −0.39 kcal mol−1 methanol.

Figure 3: Excess enthalpy (ΔH) for sinapic acid in aqueous solvents, computed using

COSMO-RS with percentage contribution denoted

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As these ΔH values were calculated based on the σ-profiles shown in Fig. 2, it is not

surprising that the DESs have much greater magnitude of ΔH compared to methanol,

due to more complementary σ-profiles. The contributions from each type of molecular

interactions are also shown in Fig. 3. H-bonding represented at least 75% of the

molecular interaction for all systems, collaborating with the interpretations from the σ-

profiles. From Fig. 3 it appears that the choice of the DES’s HBD greatly affects the

magnitude of maximum ΔH, which collaborates with the experimental results in Fig 1.

For example, ChCl: Gly (1: 1) has a much higher absolute value of ΔH relative to ChCl:

GC (1: 1), with greater magnitude of attractive ΔH (misfit) and ΔH (H-bonding) by 54.6

% and 43.63 %, respectively. Meanwhile, repulsive ΔH (VDW) was also increased by

42.50 % when glucose was used instead of glycerol which reduced solubility. This was

likely due to the larger size of the glucose molecule. Therefore, judging from

comparison of ΔH, glycerol appears to be a better choice. This is because aside from

more types of interaction having an effect, the aforementioned solvent-solvent

interaction could also play a role.

When comparing Fig. 1 and Fig. 3, it can be seen that the thermodynamics alone are

not sufficient to explain the findings from RSM and pure sinapic acids. This is because

the kinetics also play an important role. The same H-bonding behavior displayed by

DESs also leads to very high viscosities (Ozturk et al., 2018b). However, the viscosities

of the aqueous DESs was much lower than viscosities of pure DESs due to the addition

of water. For example, at 40 °C the viscosities of pure ChCl: GC (2: 1), ChCl: Gly (1: 2)

and ChCl: EG (1:2) are approximately 4000, 143 and 22 mPa s, respectively (Hayyan et

al., 2013; Ozturk et al., 2018b). While for the aqueous DES, this was 18.2, 8.60 and

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4.80 mPa s for ChCl: GC (2: 1), ChCl: Gly (1: 2) and ChCl: EG (1:2), respectively. This

allows for better mass transfer when aqueous DESs was utilized. The relative viscosity

of the DESs reflects the polarity of the HBD, with glucose being the most polar, as seen

earlier in the σ-profiles.

Even though the magnitude of viscosities had decreased, aqueous DESs demonstrate

the same trends as the pure DESs from literature. As with pure there was a significant

decrease in viscosity with increasing temperature DESs (Hayyan et al., 2013; Ozturk et

al., 2018b). This confirms that mass transfer improves with temperature, which

contributed to the increasing sinapic acid yields in Fig. 1a. The effect of DES

composition was also the same as literature, with significant difference in the viscosity

of glucose based DESs (Hayyan et al., 2013), and less so for ChCl: Gly and ChCl: EG

(Ozturk et al., 2018b). Glucose-based DESs remained the most viscous, ranging from

52.5 mPa s for ChCl: GC (1: 1) at 40 °C to 9.4 mPa s for ChCl: GC (2: 1) at 60 °C. The

lowest value for ChCl: GC is still higher than the most viscous ChCl: Gly at 40 °C, which

was at 8.9 mPa s. This shows that ChCl: Gly and ChCl: EG were able to out preformed

ChCl: GC due to their more preferable kinetics. The viscosity of ChCl: EG could be up

to 53 % lower than ChCl: Gly at the same condition. This could explain the similar

sinapic acid yields between the two types, with the thermodynamically favorability of

ChCl: Gly being counteract by the better mass transfer of ChCl: EG.

Viscosity differences also explains the significant difference between the

performances of DESs and methanol on pure sinapic acid. The dissolutions were

assessed at 30 °C leading to viscosity being much more significant. The extremely low

viscosity for the aqueous methanol, coupled with very high viscosities of the DESs

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being the likely cause. This is in addition to the effects caused by the complex matrix

within the RSM, which was not present for pure sinapic acid.

While DESs are more thermodynamically favorable for sinapic acid extraction, the

correct operating conditions are necessary in order to maximize phenolic yields. It was

demonstrated that EG and glycerol-based DESs out preformed methanol for sinapic

acid extraction from RSM, providing a better choice in terms of sustainability, safety,

efficacy and, importantly, suitability with downstream fermentation of post-extraction

RSM.

3.1.4 Phenolic extraction scale-up

While both glycerol and EG-based DESs displayed high sinapic acid yields on RSM,

for large scale extraction ChCl: Gly was chosen as the better candidate because of

better performance (Fig. 1a) and favorable thermodynamic behavior as supported by

COSMO-RS calculations (Fig. 3). Methanol extraction was also conducted for

comparison and to also assess the possibility of residual methanol in the hydrolysate

having a negative impact on the fermentation. To provide sufficient post-extraction RSM

for the bioreactor, the scalability of the phenolic extraction process was evaluated. It

was found that mass transfer was improved as a result of the scale-up. This was likely

because the ratio between the agitator length and height of the liquid column was

increased, from 0.12 to 0.58, resulting in more effective suspension of RSM. Hence,

yields for both ChCl: Gly (1: 1) and methanol were improved by 6.5 and 12.4 % wt.,

respectively, as shown in table 2.

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Table 2: Sinapic acid extraction scale up

Extraction method Extracted sinapic acid

(mg g−1 RSM)

Yield

(% wt.)

10 ml 2.5 l 10 ml 2.5 l

ChCl: Gly (1: 1) (aq) 7.63 8.21 85.0 91.5

Methanol (aq) 4.98 6.10 55.6 68.0

3.2 Bioreactor fermentation of P. putida on RSM hydrolysate

P. putida was successfully cultivated on RSM hydrolysate at 2 l scale. As shown in

Fig. 4, growth was achieved for all RSM hydrolysates, whether or not the RSM had

undergone extraction. Specific growth rates from RSM hydrolysate range from 0.15 to

0.19 h−1 (Table 3). This is slightly slower than the control, at 0.30 h−1. However, these

growth rates are in the expected range, with previous studies reporting growth rates

ranging from 0.2 to 0.4 h−1 (Hartmann et al., 2006; Le Meur et al., 2012; Meijnen et al.,

2008). There appeared to be negligible inhibitory effect of phenolics in the hydrolysate

produced from untreated RSM. This could be due to natural resistance of P. putida

KT2440, but more likely because as a result of the solid loading used during hydrolysis,

the concentrations of phenolics were too low to have an impact. Similar bacterial growth

was also achieved when RSM was treated with methanol, which was removed by

vacuum drying.

RSM was shown to have adequate concentrations of carbon and nitrogen for healthy

growth of P. putida. The large amount of nitrogen from untreated RSM, up to 3.6 g l−1

(Fig. 4), results from the presence of native proteins within the RSM. NaOH extraction of

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the proteins resulted in a reduction of approximately 50% of the total nitrogen

concentration, as seen when comparing Fig. 4b, 4c and 4d. Even though there has

been recent application of ChCl: Gly for protein extraction (dos Santos et al., 2018).

Figure 4: Time course of P. putida fermentation a) synthetic media, b) Hydrolysate from

untreated RSM, c) Hydrolysate from RSM treated with ChCl: Gly (1:1) (aq), d)

Hydrolysate from RSM treated with MeOH (aq)

As this is a proof-of-concept study with a focus on phenolics, the method for protein

extraction was chosen based on the works by Klockeman et al., who achieved a 99%

extraction efficiency on defatted canola meal (Klockeman et al., 1997). The reduced

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extraction effectiveness for proteins was due to the change in mass transfer as a result

of scale up.

Due to the abundance of nitrogen, very high levels of growth were achieved on the

RSM hydrolysates compared to the control, with a maximum cell dried weight (CDW) of

9.05 g l−1 for the RSM hydrolysate compared to the 2.55 g l−1 for the synthetic media

culture, where growth stopped after 9 hours due to nitrogen limitation. The cell

concentrations for the culture grown on the other RSM hydrolysates were lower at 4.89

and 4.54 g l−1 for DES and methanol extracted RSM, respectively. This could be due to

the lower initial concentrations of sugars, especially glucose, relative to both the

untreated RSM hydrolysate and the control. This was caused by the increase in particle

size of the RSM from aggregation during the drying process, resulting in a reduced

surface area to volume ratio during hydrolysis. The hydrolysate produced from extracted

RSM would also contain more salts compared to untreated RSM, due to residual NaOH

from protein extraction. Higher salt concentration could have resulted in lower viable cell

concentration due to osmotic stress (Bojanovič et al., 2017).

In all of the cultures PHAs were produced, consisting of 8-carbon

polyhydroxyoctanoates and 10-carbon polyhydroxydecanoates, which were consistent

with PHA produced from unrelated carbon sources via fatty acid de novo synthesis

(Huijberts et al., 1992). PHA is produced naturally due to its role as an intracellular

organelle as well as for energy storage (Huijberts et al., 1992; Prieto et al., 2016). While

high growth was achieved on the RSM hydrolysates, there was very little PHA

accumulation, as seen from table 3 where PHA accumulation was not above 0.5 %

CDW due to the abundance of nitrogen.

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Table 3: Performance of P. putida bioreactor fermentations

Experiment µ (h−1)

Max

OD600

Max CDW

(g l−1)

Max

PHA

(g l−1)

Max

PHA

(% CDW)

Control 0.30 6.71 2.55 0.08 3.08

Native 0.15 16.50 9.05 0.02 0.30

DES & NaOH 0.13 8.37 4.89 0.01 0.35

MeOH & NaOH 0.19 9.72 4.54 0.02 0.41

However, in the control culture where nitrogen was depleted by 24 hours (Fig. 4a)

there was a sharp increase of both 8-carbon and 10-carbon PHAs after nitrogen

limitation occurred. The amount of PHA accumulated reached 3.08 % of the CDW

before glucose was depleted. For the cultures grown on RSM hydrolysates,

approximately 0.35 to 0.4 g l−1 of nitrogen was consumed, while the control only

contained an initial concentration of 0.2 g l−1. If nitrogen limitation was to be induced in

the RSM cultures, it is expected that significant PHA accumulation would occur. This

would require a carbon to nitrogen ratio of at least 30: 1. Media from both post-

extraction RSMs have a carbon to nitrogen ratio of approximately 2: 1. Therefore, it is

clear that an improvement in protein extraction is essential for the RSM hydrolysate to

become a viable media for PHA production. This is in addition to the inherent value of

the proteins which incentivize their extraction.

It is also interesting to note that concentrations of xylose, galactose and arabinose

only began to decrease significantly after glucose had been depleted (Fig. 4). This could

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be due to carbon catabolite repression (CCR), where glucose is preferentially

metabolized in the presence of multiple saccharides.

However, there is limited understanding on CCR in Pseudomonas species, especially

between saccharides. This is because Pseudomonas preferred carbon sources are not

saccharides such as glucose, but rather organic acids, which has been the focus of

CCR studies within the literature (Rojo, 2010). Nevertheless, catabolism of the other

sugars still contributed to cell growth as shown in Fig. 4b. As growth in the post-

extraction RSMs were limited, catabolism of the other sugars contributed to other

cellular processes instead Fig. 4c and 4d.

Despite P. putida being able to metabolize saccharides other than glucose, the

microorganism was unable to produce PHA even when other saccharides were in

abundance, as shown by Fig. 4 a). In fact, PHA content decreased after glucose

depletion, along with the other sugars. This again could be a result of CCR. For the

purpose of maximizing PHA production, a high glucose concentration must be

maintained, either through increasing the initial concentration or implementation of

batch-feeding strategies. From this observation, PHAs yield coefficients were also

defined with respect to glucose, Y PHAGC

. Since PHA accumulation peaked at the beginning

of the stationary phase (Fig. 4), the yield coefficients at this point were used for

comparison. As seen from table 4, yield coefficients for all cultures range from 0.006 to

0.020 g g−1, as the majority of the carbon was used for cell biomass. The RSM cultures

in fact had similar conversion rate as the control. This further suggests that if nitrogen

limitation were to be induced, higher PHA accumulation can be achieved when grown

on RSM.

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Table 4: Yield coefficients of P. putida bioreactor fermentations at the beginning of the

stationary phase

Yields coefficients (g g−1)

ExperimentY X

C

Y XC+N

Y PHAGC

Control 0.32 0.29 0.013

Native 0.94 0.32 0.006

DES + NaOH 2.59 0.68 0.020

MeOH + NaOH 1.43 0.48 0.013

Y XC: Yield coefficient for residual cell weight w.r.t. carbon sources;

Y XC+N

: Yield coefficient for residual cell weight w.r.t. carbon and nitrogen sources;

Y PHAGC

: Yield coefficient for PHA w.r.t. glucose;

Cell yield with respect to carbon sources, Y XC, for the control culture was at 0.32

(Table 4). This is similar to the growth of P. putida on synthetic media with sugars as a

carbon source found in literature. For example, La Meur et al. found that their

recombinant P. putida KT2440 had a yield of 0.5 g g−1 when grown on xylose (Le Meur

et al., 2012). On the other hand, yields from the RSM hydrolysate were extremely high,

reaching as high as 2.59 g g−1 for DES extracted RSM. This does concur with higher

growth seen on RSM hydrolysates. However, as the RSM yields are much higher than

1, it is likely that the carbon is not coming from the sugar alone. This difference could be

due to residual DES components acting as additional carbon sources. However, this

does not account for the hydrolysate produced from the methanol extracted or the

native RSMs.

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The additional carbon more likely came from the protein derivatives in the

hydrolysates. The high temperature and low pH conditions during acid hydrolysis could

have certainly induced protein hydrolysis (Williams, 2003). Additionally, catabolism of

both d- and l-amino acids has been reported on the strain of P. putida used (KT2440)

(Radkov and Moe, 2013), some of which are present in relatively large amounts in

RSM, such as lysine, phenylalanine and arginine (Tzeng et al., 1988). This corroborates

with the fact that the difference in Y XC between the hydrolysates and control was due to

the lower amount of sugars being consumed, since as stated previously, all cultures had

similar specific growth rates. Thus, it was more accurate to access the fermentation’s

performance based on the yield with respect to both the sugars and proteins, Y XC+N

. The

difference between Y XC+N

of the different fermentations is much lower compared to Y XC.

RSM hydrolysates have Y XC+N

only ranging from 0.32 to 0.68 g g−1, while Y XC+N

for the

control culture was also comparable at 0.29 g g−1. Y XC+N

values were also of similar

magnitude to yields coefficient expected from literature, such as the aforementioned

work by Le Meur et al.(2012). Davis et al. (2015) also reported a yield coefficient of 0.45

g g−1 for P. putida on glucose. These similar yield coefficients demonstrate that RSM

hydrolysate can act as an effective growth media for P. putida fermentation.

4. Conclusions

This study is the first to demonstrate an integrated process for DES extraction of high

value components followed by biopolymer production from RSM. DESs significantly

outperformed methanol for phenolic extractions, up to 85.7 % for ChCl: Gly. Selecting

appropriate DESs components can maximize extraction yield by optimizing the H-

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bonding interactions and DES viscosity. Successful scale-up of the extraction was

achieved, allowing for bioreactor fermentations of residue-derived hydrolysates.

Extremely high growth was achieved on the hydrolysates, yielding CDW of up to 4.89 g

l−1, with growth rates comparable to those obtained using synthetic media. 8-carbon and

10-carbon PHAs were also successfully synthesized using RSM hydrolysates.

Acknowledgements

The authors would like to thank Mrs. Carole Webb and the School of Chemistry, The

University of Manchester for their valuable assistance on gas and liquid

chromatography.

Appendix A. Supplementary data

Supplementary data provided with the online version of paper.

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