In Situ Lignin Bioconversion Promotes Complete Carbohydrate

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In-situ lignin bioconversion promotes complete carbohydrate conversion of rice straw by Cupriavidus basilensis B-8 Mengying Si, Xu Yan, Mingren Liu, Meiqing Shi, Zhongren Wang, Sheng Wang, Jin Zhang, Congjie Gao, Liyuan Chai, and Yan Shi ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.8b01336 • Publication Date (Web): 01 May 2018 Downloaded from http://pubs.acs.org on May 4, 2018

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ACS Sustainable Chemistry & Engineering

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In-situ lignin bioconversion promotes complete carbohydrate conversion of rice

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straw by Cupriavidus basilensis B-8

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Mengying Si, † Xu Yan, †, ‡ Mingren Liu, † Meiqing Shi, †,‡ Zhongren Wang, † Sheng

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Wang, ‡ Jin Zhang, § Congjie Gao, †, ‡,∥ Liyuan Chai, †, ‡ Yan Shi*, †, ‡

5

†

6

China

7

‡

8

Metal Pollution, Changsha 410083, China.

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§

School of Metallurgy and Environment, Central South University, Changsha 410083,

Chinese National Engineering Research Centre for Control & Treatment of Heavy

College of Environmental Science and Engineering, Hunan University, Changsha

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410082, P.R. China.

11

∥

12

Technology, Hangzhou 310014, PR China

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* E-mail: [email protected] (Y. Shi); Fax: +86-0731-88710171; Tel:

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+86-0731-88830875

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M. Si, X. Yan, M. Liu, M. Shi, Z. Wang, S. Wang, L. Chai, Y. Shi, Mailing address:

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No.932 South Lushan Road, Changsha Hunan 410083, P.R. China

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J. Zhang, Mailing address: No. 2 South Lushan Road, Changsha Hunan 410082, P.R.

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China

19

C. Gao, Mailing address: No.18 Chaowang Road, Hangzhou Zhejiang 310014, P.R.

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China

Water Treatment Technology Development Center, Zhejiang University of

1

ACS Paragon Plus Environment

ACS Sustainable Chemistry & Engineering 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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ABSTRACT

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The valorization of lignocellulose encounters both opportunities and challenges

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as the lignocellulose is an abundant and intrinsically heterogeneous natural source.

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However, the successful design of an integrated process for complete carbon

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utilization of lignocellulose is limited. A classic base-catalyzed pretreatment strategy

26

and a natural Cupriavidus strain with the capacity of lignin degradation and

27

polyhydroxyalkanoates (PHA) biosynthesis were selected to establish a fundamental

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and functional module necessary to enable a new platform for lignocellulose

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pretreatment and waste carbon conversion. The in-situ bioconversion was first

30

introduced to the pretreatment. Specifically, selectively cleaving insensitive C–C

31

bonds (e.g., β-5) of lignin via a “washing” mechanism, Cupriavidus basilensis B-8

32

promoted the digestibility of the rice straw to realize almost complete carbohydrate

33

conversion, yielding 984.2 mg g-1 of reducing sugar when combined with alkaline

34

pretreatment. A demonstrated concentration of PHA (482.7 mg L-1) was obtained

35

from the conversion of the removed lignin in ligninolytic bacteria. The integrated

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molecular conversion mechanisms of lignin in bacteria were further elucidated. Our

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work provides a novel perspective for biorefinery design.

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KEYWORDS: Cupriavidus basilensis B-8, in-situ bioconversion, delignification,

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lignocellulose pretreatment, polyhydroxyalkanoates, carbohydrate conversion

2


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INTRODUCTION

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Lignocellulose is composed primarily of carbohydrate polymers (cellulose

42

and hemicellulose) and a heterogeneous matrix of phenolic polymers known as

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lignin.1 Lignin offers lignocellulose recalcitrance to inhibit both enzymatic

44

saccharification and the fermentation of carbohydrates. Further in the

45

pretreatment, lignin depolymerization has recently evolved as a key step to

46

enable saccharification of cellulose.2 Lignin, in the nature, is an amorphous and

47

complex three-dimensional heteropolymer composed of cross-linked phenyl

48

propane units via a variety of ether and carbon–carbon (C–C) bonds, thereby

49

forming a physical barrier to protect biomass from microbial attacks.2-3 The

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typical pretreatments (e.g., dilute alkali/acid pretreatments) mainly cleave

51

β-O-4 ethers to depolymerize lignin,4 leaving the insensitive C–C bonds as a

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restrictive factor hindering the achievement of complete carbohydrate

53

conversion.5 The 5-5 and β-5 dimeric lignin model compounds could be

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selectively oxidized by laccase,6 which was considered as a major ligninolytic

55

enzyme in microorganisms. Hence, it shows a great potential to employ the

56

ligninolytic strains to conquer the restrictive factors of delignification for

57

advanced pretreatment.

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As compared to chemocatalysis, biocatalysis has attracted increasing

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attention as an environmentally friendly and low cost method.7-8 To date,

3



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biocatalysis in pretreatments has mainly focused on white rot fungi harboring

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complex ligninolytic enzymatic systems, which were thought to depend in part

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on extracellular heme-dependent peroxidases, copper-dependent laccases, and

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other oxidases to depolymerize lignin. However, a commercial biocatalytic

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process of lignin depolymerization has yet been reported, partially due to their

65

poor stability in industrial processes and practicality challenges, specifically in

66

fungal protein expression and genetic manipulation.9-10 Therefore, lignin

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utilization by bacterial species with more rapid growth and easier genetic

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manipulation has been considered as a promising biorefining method.

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In addition, lignin represents an untapped resource for the production of

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biofuels and valuable chemicals, as it is the most abundant natural source of

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aromatic compounds in the biosphere.11 Unfortunately, the lignin fraction of

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plant cells has been mainly employed for heat and power via combustion in the

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pulp and paper industry.12-13 Recent studies on the proposed lignocellulosic

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valorization have concluded that lignin depolymerization and conversion assists

75

in the utilization of biomass to useful products due to co-valorization and the

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presence of new revenues beyond carbohydrates.4 Therefore, one of the greatest

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challenges in biorefining is the engineering of lignin structures to not only

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remove or modify lignin to reduce lignocellulose recalcitrance but also to

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enable lignin valorization.14 Recent research has focused on bacterial

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conversion of lignin to lipids,15 adipic acid,16 and polyhydroxyalkanoate 4


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(PHA).17 Hence, it is feasible to design an integrated biocatalytic process to

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simultaneously achieve the high sugar yield and lignin valorization.

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In our previous research, the ability of Cupriavidus basilensis B-8

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(hereafter B-8) to selectively degrade Kraft lignin without the presence of a

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co-substrate was validated. The conversion of Kraft lignin to PHA was also

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confirmed.18 Herein, rice straw (RS), the largest available and deserted biomass in

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the world,19 was employed as the feedstock. B-8 was employed not only to

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in-situ bioconvert the natural lignin in RS, but also to enhance the enzymatic

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digestibility. To our knowledge, it is the first study that realized the in-situ

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bioconversion of lignin during the pretreatment process. Moreover, the mechanism

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of lignin conversion was investigated. Our study provides a new platform for

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lignocellulose pretreatment and waste carbon conversion.

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EXPERIMENTAL SECTION

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Lignocellulose, strain and culture medium. The rice straw (RS)

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obtained from Shandong, China was ground into powder and air-dried. The

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air-dried RS powder was then sifted using a 60-mesh griddle and used as the

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pretreatment feedstock. The Cupriavidus basilensis B-8 (CGMCC No. 4240,

98

hereafter B-8) was overnight cultured on a rotary shaker at 30 °C with a speed of 150

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rpm in a Luria-Bertani broth medium, and then diluted five-fold in Luria-Bertani

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broth medium with continued growth for 2 h to achieve the logarithmic growth phase.

101

The obtained bacterial culture was used as the seed culture for the biological 5



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treatment.

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Bio- and chemocatalysis for the valorization of the rice straw. 2 g

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untreated RS was soaked in a 20-mL NaOH solution with different

105

concentrations (0.5%, 1%, and 2%) and then statically treated at 121 °C at

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different times (15 min, 30 min, and 60 min). The NaOH-treated RS sample

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was collected and thoroughly washed with de-ionized water, then dried at 50°C

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for further pretreatment. To obtain a lignin-rich stream for upgrading, alkaline

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pretreated liquor (APL) was collected during the chemical process. The B-8

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based biocatalysis was conducted as follows: 10 mL seed culture was

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centrifuged to harvest the bacterial cells. The collected cells were then

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inoculated into 100 mL sterile mineral salt medium

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NaOH-treated RS. To support growth, APL was adjusted to a pH of 7.0 by

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slowly wadding 5 M H2SO4 and was supplemented with mineral salt. A

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subsequent treatment was similarly employed as the process mentioned above.

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All the biocatalysis processes were performed in a rotary shaker at 30°C with a

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speed of 150 rpm.

20

containing 1 g

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Enzymatic hydrolysis. Commercial cellulase (Cellic CTec2, Novozymes,

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Bagsværd, Denmark) was used for the enzymatic hydrolysis. A typical

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hydrolysis mixture consisted of 0.5 g RS sample, 20 mL of the 50-Mm citric

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acid buffer (pH 4.8), which was supplemented with cellulase (12 filter paper

122

units (FPU) g-1), antibiotic (40 µg mL-1), and cycloheximide (30 µg mL-1) to 6


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prevent microbial contamination.21 The mixture was incubated at 50°C in a

124

rotary shaker at 110 rpm for 24 h. Samples were then collected and centrifuged

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for sugar analysis. The reducing sugar was measured by the 3,5-dinitrosalicylic

126

acid assay.22

127

Characterization of the biomass. Different treated RS samples were

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collected for the composition and physicochemical analyses. The chemical

129

compositions were measured according to the method presented by Teramoto.23

130

Solid gold-coated RS samples were observed using a scanning electron

131

microscope (SEM, JSM-IT300LA, JEOL, Japan). Atomic force microscopy

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(AFM) imaging was performed in tapping mode on a NanoManTM VS +

133

MultiMode V scanning probe microscope (Veeco Company, USA).24 The

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crystallinity of the RS were analyzed using a TTR III X-ray diffractometer

135

(Rigaku, Japan).25

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The lignin fraction was extracted from the treated and untreated RS as well

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as APL.26 The molecular weight of lignin was then measured by gel permeation

138

chromatography (GPC, waters 1515, Waters Company, USA).17 A 1760X

139

Fourier Transform Infrared (FTIR) spectrometer (PerkinElmer, Shanghai, China)

140

and a Bruker Avance 500 MHz spectrometer (Bruker GmbH, Karlsruhe,

141

Germany)27-28 were employed to analyze the structural changes.

142

Polyhydroxyalkanoate (PHA) production by B-8 cultivation. The

143

nitrogen (N) concentration was strictly controlled during cultivation. 20 mL of 7



144

the collected cells were inoculated into a 100 mL sterile N-limit mineral salt

145

medium ((NH4)2SO4 concentration: 30 mg L-1) containing an excess of

146

NaOH-treated RS or APL. The PHA accumulation was monitored by

147

fluorescence microscopy. To determine the cell dry weight, 200 mL of culture

148

was centrifuged, washed in 10% PBS, recentrifuged, and lyophilized. The

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PHAs were recovered by accelerated solvent extraction with methanol and

150

chloroform (presented in Supporting Information).

151

Characterizations of the PHA. The number and weight average

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molecular weights (Mn and Mw) of the standard and sample PHAs were

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measured using a PL-GPC 120 (Polymer Laboratories). The thermal properties

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of the PHAs were analyzed by differential scanning calorimetry (DSC) and

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thermal gravimetric analysis (TGA).29 Gas chromatography analysis was

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performed using a GC QP2010 MS (Shimadzu, Kyoto, Japan). The samples

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were prepared by methanolysis. The major functional groups present in the

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purified PHA polymer were detected by FTIR spectrometry. Furthermore, the

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microchemical structure of the PHAs was investigated using a Bruker Avance

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500 MHz spectrometer at 500 MHz for 1H analysis.

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Transcriptional analysis. The B-8 cultures were grown using fructose,

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NaOH-treated RS, and APL as the carbon sources. The cells were harvested at

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the end of their exponential growth.30 The RNA preparation, library

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construction, and sequencing on a BGISEQ-500 was performed at the Beijing 8


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31

165

Genomics Institute.

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remove the reads with adaptors, reads with more than 10% unknown bases, and

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low-quality reads. The purified reads were obtained and stored in FASTQ

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format.31 HISAT32 was employed to map the clean reads to the genome. The

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gene expression levels were quantified by a software package called RSEM.33

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The NOISeq method was employed to screen the differentially expressed genes

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into two groups.

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up-regulated genes were analyzed to illustrate the mechanism.

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RESULTS AND DISCUSSION

34

All the generated raw sequencing reads were filtered to

After filtering (≥ 2-fold change, adjusted p ≤ 0.001), the

174

Potential biocatalysis of Cupriavidus basilensis B-8 (hereafter B-8) for

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lignocellulose valorization. Although the enzymology of bacterial lignin

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degradation has not been as thoroughly investigated as that of fungi, indications

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have suggested the evolution of bacteria into their own enzymatic systems for

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lignin degradation.35 We previously observed the activities of laccase and

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manganese peroxidase in B-8.20 Moreover, the presence of the Fenton reaction

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in the depolymerization of lignin by B-8 is possible.20 Herein, we performed a

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comprehensive and systematic whole genomic analysis on the lignin-related

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aromatic compounds degradation pathways based on previously obtained

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genomic data of B-8.20 A significant number of genes encoding determinants

184

that constructed five pathways involved in lignin degradation were identified 9



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185

(Figure S1). They included two branches of the β-ketoadipate pathway (cat

186

genes and pca genes), phenol pathways (mml genes and phl genes),

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phenylacetyl-CoA ring-cleavage pathways (paa genes), the gentisate pathway

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(mhb genes), and the 2,3-dihydroxyphenylpropionate catabolic pathway (mhp

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genes).

190

acetyltransferase),

191

(poly(3-hydroxyalkanoate) synthetase), and phaR (polyhydroxyalkanoate

192

synthesis

193

polyhydroxyalkanoate (PHA), was also identified.

The

pha

repressor)

gene

cluster,

phaB

and

which

includes

(acetoacetyl-CoA

is

responsible

for

phaA

(acetyl-CoA

reductase),

the

phaC

biosynthesis

of

194

Carbohydrates such as cellulose and hemicellulose account for about 75%

195

of lignocellulosic dry weight. All the carbohydrates are expected to be

196

saccharified for ethanol production. To enhance the enzymatic digestibility of

197

the RS, and avoid carbohydrate loss, the potential of the selective lignin

198

removal in the bacteria is needed. Hence, different types of carbohydrates,

199

including glucose, fructose, sucrose, galactose, xylose, arabinose, mannose,

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lactose, maltose and pullulan were selected as the sole carbon sources for B-8

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in the present study. According to Fig. S2, B-8 only used fructose for growth

202

during the first 3 days, and hardly used any of the typical monosaccharide

203

components of cellulose (glucose) and hemicellulose (xylose, galactose,

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arabinose, mannose, lactose, and maltose) as the sole carbon source (Figure S2).

205

The similar results were generated even after an extended incubation time of 10


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one week. Therefore, B-8 exhibited outstanding potential to selectively

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depolymerize or degrade lignin component for the deep pretreatment of

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lignocellulose and PHA synthesis.

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Biochemical pretreatment significantly promotes the enzymatic hydrolysis

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of the RS. To evaluate the efficiency of the pretreatments, the RS residues were

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hydrolyzed with commercially available cellulase CTec2 for 24 h. The results are

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presented in Figure 1a. Obviously, the NaOH pretreatment significantly improved

213

the enzymatic digestibility of the RS. The saccharification of the untreated samples

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for 24 h at a cellulase loading of 12 FPU g−1 resulted in only 91.6 mg g-1 reducing

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sugar. Following NaOH pretreatment, the reducing sugar exhibited an improved yield

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of 350.9-849.2 mg g-1. These results can be attributed to the nucleophilic cleavage of

217

the lignin phenolic alkyl-aryl ethers and the promotion of the solvation of lignin into

218

its aqueous phase during the alkaline pretreatment,

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fractionation of 37.4%–84.6% lignin (Figure 1b) to the alkaline pretreated liquor

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(APL). The maximum sugar yield of 849.2 mg g-1 was obtained under the relative

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severe condition (2% NaOH, 60min, 121°C). Though the severe pretreatment

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removed 84.6% lignin, the residual lignin still presented a significant challenge for the

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enzymatic hydrolysis. Sequentially, biopretreatment by B-8 was employed. After B-8

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pretreatment, more lignin (49.2%–88.5%) were removed, leaving purer carbohydrates

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(maximum 91.2%) with lighter in color appearance (Figure 1b). And the enzymatic

226

digestibility of RS was significantly improved (13.5%–32.6%, compared to sole

4

which was convinced by the

11



227

NaOH pretreatment). It's noted that B-8 promoted the digestibility of the RS to

228

achieve the almost complete carbohydrate conversion with the sugar yield of 982.6

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mg g-1. Particularly, this demonstrated efficiency of pretreatment showed a superiority

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as compared with other researches (Table S1).8,

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employed linear fitting to test the correlation between the lignin content and the sugar

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yield (Figure 1c), of which a strong negative correlation (R2=0.829) was observed,

233

thereby indicating that the decrease in the lignin content positively affected the

234

enzymatic hydrolysis of the RS samples. As one of the most key barriers, lignin not

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only physically limited the activated cellulose to the accessible surface but also

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unproductively bind itself to the enzymes through the functional groups such as the

237

phenolic hydroxyl groups. 40-41 These results suggested the important role of B-8 in

238

removing the residual lignin in biomass after alkaline pretreatment to obtain purer

239

carbohydrates and better digestibility of biomass.

36-39

Moreover, the present study

240

To explore these mechanisms, samples from the co-pretreatment of RS under the

241

optimum condition was selected for further characterization. SEM images (Figure 2a)

242

presented that untreated RS always had a regular and tough structure with a smooth

243

and compact surface, whereas the NaOH treatment exhibited digestion holes or pits.

244

Further bacterial treatment of C. basilensis B-8 verified its outstanding potential to

245

specifically metabolize lignin fraction as RS appeared to be peeled off individual

246

5-μm fibrous structures from the surface by specific delignification. The microscopic

247

morphology characterization of the untreated and pretreated RS samples allowed us to 12


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propose a model system, “washing” mechanism, as elucidated in Figure 2b.

249

Specifically, the NaOH solution catalyzed the dissolution of lignin to the aqueous

250

phase and retained the holocellulose framework of the biomass, thereby retaining the

251

lignin fragment in a fashion similar to remain blots on the clothing fibers (framework

252

of RS). Surprisingly, B-8 acted as a scavenger that specifically “washed off” the

253

residual lignin fragment between the interlaced fibrous structures. This hypothesis

254

was verified by the nearly invariable cellulose crystallinity results following various

255

pretreatments, which exhibits minimal cellulose fraction disruption during the

256

pretreatments (Figure S3).

257

To further verify the “washing” mechanism, a nanoscale visualization of RS

258

morphology was analyzed by the AFM images (Figure 2c-e). Generally, the

259

accessibility of cellulose, which depends on the opened cellulose surface area and the

260

material porosity

261

increased porosity improves the access of cellulase to cellulose. In general, the color

262

in the height maps represents the surface roughness of RS. The RS height maps

263

exhibited a lighter color in Figure 2c, thereby indicating that the initially smooth

264

untreated RS surface (Ra: 8.47 nm, Rmax: 93.4 nm) gradually roughened (Ra: 12.9 nm,

265

Rmax: 179 nm and then Ra: 25.6 nm, Rmax: 274 nm). It might be attributed to the

266

chemical process and biocatalysis-induced increase in RS substrate porosity to expose

267

more surface area.43 In addition, the AFM tip adhered more strongly to the

268

hydrophilic area that hence appeared lighter in color in the phase diagram.44 The same

41

is assumed to play an important role in hydrolysis.42 Thus, the

13



269

area in both height map and phase diagram presented uniform changes in color.

270

Specifically, the exposed surface area from the treatment (lighter area in the height

271

map) exhibited more hydrophilic cellulose that appeared lighter in the phase diagram.

272

Based on the SEM and AFM analyses, B-8 can selectively “wash off” the

273

hydrophobic lignin “blot” to expose the activated cellulose surface and decrease the

274

unproductive binding of enzymes on lignin, thereby increasing cellulose accessibility.

275

Moreover, GPC, FTIR, and 2D NMR analyses were employed to explore the

276

chemical mechanism of lignin degradation involved in the “washing” mechanism.

277

Firstly, to understand variations in the degree of lignin polymerization in the RS

278

during pretreatment, GPC analysis was performed to determine the weight-average

279

(Mw) and number average (Mn) molecular weights as well as polydispersity (PDI,

280

Mw/Mn) of the isolated lignin fractions (Table 1, Figure S4). The Mw of untreated

281

lignin and NaOH pretreated lignin was 3105 Da and 1399 Da, respectively, implying

282

the sensitive linkages in lignin were cleaved by NaOH pretreatment. After B-8

283

pretreatment, the Mw of lignin were further decreased to 868 Da, indicating the

284

promotion of lignin depolymerization by B-8. According to FTIR results (Figure S5

285

and Table S2), the bands at 1513 cm-1 and 1420 cm-1, which correspond to aromatic

286

skeletal vibrations and the C-H deformation, respectively, combined with the aromatic

287

ring vibration at 1460 cm-1 were significantly reduced, also indicating that the lignin

288

units were gradually removed following each pretreatment step. Specifically, the

289

syringyl (S) unit breathing with C=O stretching at 1329 cm-1 was not observed in the 14


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290

RS following the NaOH treatment, showing that most S units in lignin were easily

291

removed by the alkali solution. Moreover, the intensity of the C–H out-of-plane units

292

in positions 2, 5, and 6 of the guaiacyl (G) rings at 858 cm−1 was weaker both in

293

NaOH and Co-pretreatment lignin, indicating that G-unit was the common target of

294

NaOH and B-8 pretreatment.

295

The 2D-HSQC NMR analysis generates important structural information on the

296

lignin macromolecule such as the monolignol compositions (including S/G ratios) and

297

the relative abundance of the inter-unit linkages.45 The S/G ratio dramatically

298

decreased following NaOH treatment from 1.28 to 0.82 (Figure 3 and Table 1),

299

thereby indicating the easier removal of S units under the alkaline condition, which is

300

consistent with the FTIR spectra results. Meanwhile, the p-hydroxyphenyl (H) units

301

disappeared during the chemical process. Differently, the S/G ratio increased to 2.09

302

after B-8 pretreatment, thereby indicating that B-8 mainly metabolized the G units.

303

The relative-quantification of the lignin fractions by the 2D-HSQC NMR method

304

provides the explicit structural evolution during the pretreatments. Herein, Table 1

305

presents the changes observed in the lignin chemical bonds during pretreatment.

306

Lignin from the untreated RS was rich in β-O-4 linkage (86.3%), which exhibited no

307

obvious changes during the NaOH treatment (88.6% in NaOH-RS). Considering

308

lignin were largely (84.6%) removed by NaOH pretreatment, the unconspicuous

309

change in β-O-4 linkage content in lignin demonstrated that NaOH pretreatment 15



310

cleaved most of the β-O-4 linkages. Specifically, the signals of Aβ(G) and Aβ(S)

311

declined, thereby attributing to the depolymerization of the β-O-4 aryl ethers.

312

Meanwhile, signals associated with β-β were not detected in NaOH pretreated lignin,

313

possibly due to the removal of the S units after NaOH pretreatment.46 In terms of the

314

biopretreatment, the weaker signals associated with the phenylcoumaran (β-5-coupled)

315

structures were primarily derived from the decrease in G and H units, implying that

316

the C–C linkages (β-5) were more easily cleaved during the removal of the G units by

317

B-8. In summary, the S and H units in lignin were mainly removed via the cleavage of

318

the sensitive β-O-4 linkages during NaOH pretreatment. As a major restrictive factor

319

of lignin depolymerization, the remained C–C linkages (β-5) structure in residual

320

lignin inhibit the further delignification. Fortunately, B-8 selectively remove the G

321

units by cleaving the alkaline-resistant β-5 linkages via the "washing" mechanism to

322

promoted the digestibility of the RS to realize almost complete carbohydrate

323

conversion.

324

Biocatalysis efficiently depolymerizes the lignin in APL. Following the NaOH

325

pretreatment, the complex streams containing components with different molecular

326

weights derived from both lignin and polysaccharide were produced. Due to the high

327

level of chemical heterogeneity, these streams tended to be metastable and were

328

typically underused. In the presented integrated process, the waste stream also

329

exhibited valorization to maximize carbon utilization by biocatalysis. Given that

330

lignin is the most abundant component in the waste stream, understanding of the 16


Page 16 of 43

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331

lignin conversion mechanism by B-8 is important for the integrated process of

332

lignocellulose valorization. Herein, we first studied the bacteria-induced lignin

333

changes at the chemical level via GPC, FTIR and 2D NMR analyses. Figure S6

334

presented the Mw, Mn, and PDI results of the lignin obtained from APL, wherein

335

B-8-treated APL exhibited a smaller Mw (1444 Da) as compared to the untreated APL

336

(1705 Da), which may possibly be due to the release of small lignin fragment

337

molecular weights following with lignin depolymerization. To confirm the above

338

hypothesis, the structural changes in the lignin fractions were characterized by FTIR

339

and 2D-HSQC.

340

The aromatic region in the spectra of all the lignin fractions (Figure 4a)

341

displayed the basic lignin units (G, S, and H units). An increase in the

342

B-8-treated-APL S/G ratio from 1.28 to 3.07 indicates that the biocatalysis process

343

selectively removed lignin fragments with higher G unit contents and left S unit-rich

344

lignin in the APL. This was verified not only by the decreased signals at FA2 and FA7

345

but also by the weakened band assigned to the aromatic skeletal vibrations (G>S)

346

around 1513 cm-1 (Figure S7, Table S1). In addition, the p-coumarates and H units in

347

lignin were also depolymerized, which was certified by the decrease of the signals at

348

PCE 3, 5 and the H units.

349

Furthermore, the side-chain regions of the 2D-HSQC NMR spectra reflect the

350

types and distribution changes of the inter-unit linkages in the lignin fraction.47

351

According to Figure 4c, lignin from the untreated APL was rich in β-aryl ether units 17



352

and had modest amounts of resinol and low amounts of phenylcoumaran, respectively,

353

as well as modest levels of cinnamyl alcohol end groups. The signals at Aβ(G) and

354

Aβ(S) assigned to the β-position in β-O-4 were linked to the G and S units (A),

355

respectively. Both units disappeared during cultivation. These variations were mostly

356

attributable to the cleavage of the β-O-4 aryl ethers. Given that the β-5 structural unit

357

only stemmed from the coupling monomer of the G and p-hydroxyphenyl (H) units,48

358

a reduction in the amount of G units resulted in a decrease in the phenylcoumaran

359

(β-5-coupled) structures. In addition, the overserved resinols (β-β-coupled units) were

360

mainly raised from the dimerization of sinapyl alcohol. The structure of cinnamyl

361

alcohol end groups (I) disappeared primarily due to coniferyl alcohol (G unit)

362

depolymerization. All of the results indicate that the biocatalysis strategy of B-8

363

efficiently depolymerizes the lignin in APL.

364

PHA production and characterization. The B-8 cultures were respectively

365

inoculated in a mineral salt medium with APL and NaOH treated RS as the sole

366

carbon source. Nile red staining under fluorescence microscopy confirmed the

367

accumulation of PHA (Figure S8). Although few carbon source (only 2.88% lignin)

368

left in the NaOH-treated RS, 483.4 mg L-1 of dry cell weight was harvested, and 32.7

369

mg L-1 of PHA was accumulated. It indicated that lignin was depolymerized and

370

in-situ bioconverted to PHA by B-8. Meanwhile, a maximum dry cell weight of

371

3990.2 mg L-1 was harvested and PHA with a weight of 450.0 mg L-1 was

372

accumulated by B-8 from the APL. To evaluate the efficiency of lignin bioconversion 18


Page 18 of 43

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373

by B-8, a comparison was summarized in Table S3. Currently, Pseudomonas putida

374

KT2440 grown in the fed-batch fermentation mode has accumulated a record level

375

concentration (1.0 g L-1) of PHA using lignin stream that was obtained from the

376

combinatorial pretreatment of sulfuric acid and NaOH.49 NaOH not only removes

377

most of the lignin, but also slightly attacks cellulose and hemicellulose through

378

“peeling” reactions. While sulfuric acid mainly attacks hemicellulose and cellulose,

379

and simultaneously modifies the lignin. Therefore, the produced liquor contained

380

abundant carbon including lignin-related component with small molecular weight as

381

well as sugar and organic acids originated from cellulose and hemicellulose. However,

382

when anthraquinone was cocharged to the reactor of NaOH pretreatment to maximize

383

polysaccharide retention in the solids via the minimization of polysaccharide “peeling”

384

reactions, a maximum PHA yield of only 252 mg L-1 was obtained.17 In addition, we

385

found that P. putida KT2440 hardly grow in the medium with lignin (kraft lignin or

386

lignin extracted from RS) as sole carbon source. Moreover, a genetically engineered

387

bacterium P. putida A514 only yielded 75 mg L−1 of PHA under kraft lignin alone.50

388

Catabolized sugar or organic acids originating from polysaccharide breakage during

389

lignocellulose pretreatment will likely improve lignin conversion. This hypothesis

390

provided a new perspective for more optimal engineering of ligninolytic bacteria for

391

the valorization of lignin.

392

Analysis of the hydroxy acid monomer distribution of PHA from APL indicated

393

that the PHA polymer was primarily comprised of 2-hydroxybutyrate acid (2HB 19



394

9.9%), 3-hydroxybutyrate acid (3HB 86.5%), and 3-hydroxyvaleric acid (3HV 3.6%).

395

The RS-derived PHA was determined to be a copolymer that was mainly composed of

396

2HB (5.9%), 33HB (92.9%), and 4-hydroxyheptanoate acid (3HHp 1.2%) (Figure S9).

397

Moreover, the spectra results determined by FTIR analysis (Figure S10) and 1H-NMR

398

analysis (Figure S11) indicated the presence of typical PHA. The produced PHAs

399

(both from APL and RS) exhibited an Mw of 1694 kDa and 1160 kDa with a PDI of

400

1.58 and 1.48, respectively (Figure S12). The lower dispersity indicated the presence

401

of more homogeneous polymers. In addition, the melting (Tm) and decomposition (Td)

402

temperatures (Figure S13) of PHA from APL (167.5°C and 273.0°C, respectively) and

403

PHA from the RS residue (162.8°C and 265.4°C, respectively) exhibited excellent

404

thermostability. The characterization of the PHAs indicated that their physicochemical

405

properties were comparable to those derived from carbohydrates.51

406

Effect of lignin removal from APL and the NaOH-treated RS residue under

407

the nitrogen (N) limited condition. The polymer accumulated in B-8 during growth

408

under an N-limited condition. However, the N-limited condition negatively affects

409

bacterial growth (or biomass yield) and cell function. Therefore, the following

410

question is posed: does the stress condition (N-limitation) affect the removal of lignin

411

from APL and the biocatalysis for lignin removal to improve enzymatic hydrolysis?

412

To answer this question, an investigation on the removal of lignin from APL and the

413

RS residue following NaOH pretreatment with different bacterial cell inoculations

414

was performed. 20


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415

Herein, A280 measurements and the reduction in total phenols were used to

416

quantify the removal of lignin. 30.7% of lignin was removed based on A280 and 21.8%

417

based on the total phenols of APL with 10% of the bacterial inoculation under the

418

N-rich condition (Figure 5). However, the increase in A280 and the decrease in the

419

total phenols were all less than 10% at a 10% bacterial inoculation under the

420

N-limited condition, thereby indicating that the lignin removal decreased by at least

421

60% for bacterial growth in an N-limited culture. Interestingly, when the bacterial

422

inoculation increased to 20%, lignin removal under the N-limited condition was

423

comparable to that with 10% of bacterial inoculation under the N-rich condition. The

424

NaOH pretreatment of the RS removed 84.6% of lignin and offered a great challenge

425

for the quantification of lignin residue on the RS. Therefore, the reducing sugar

426

release was used to evaluate the effect of the RS residue on the lignin removal

427

following NaOH pretreatment by bacteria grown under the N-limited condition. No

428

obvious differences were observed between the reducing sugar release from the

429

co-treated RS for bacterial growth under the N-limited condition and from

430

NaOH-treated RS (Figure S14). However, a bacterial inoculation increases to 20%

431

resulted in a 19.8%-increase in the reducing sugar release as compared to that with

432

NaOH-pretreatment alone. This was comparable to the release reducing sugar after

433

the co-pretreatment under the N-rich condition.

434

Molecular mechanism. To further explore the insights of such consolidated

435

biomass processing, a transcriptomics analysis was performed on B-8 grown on the 21



436

APL and NaOH-treated RS (Figure 6, S15 and Table S4-10). The results (Table S4)

437

revealed the presence of an up-regulated gene encoding multicopper oxidase that was

438

negligible in the genomic search of the APL and NaOH-treated RS culture. It

439

indicated that laccase-based lignin depolymerization presented in B-8. Several

440

peroxidases were observed and identified, specifically cytochrome c peroxidase and

441

catalase/peroxidase. However, the role of these enzyme for the degradation of lignin

442

in B-8 remains unknown. The results characterize the peroxidase-based lignin

443

depolymerization as the core component of the lignin depolymerization mechanisms

444

and is certainly essential for B-8. Moreover, NADPH quinone oxidoreductase were

445

found with aryl alcohol oxidase and pyranose-2-oxidase, which supports Fenton

446

chemistry through generating extracellular H2O2 involved in lignin depolymerization.

447

Through the lignin depolymerization process, the released low molecular weight

448

compounds can be transported into the cell for complete aromatic compounds

449

catabolism. The over-expressed cat, pca, ben and hca genes (Table S5) indicate that

450

the β-ketoadipate and peripheral pathways

451

B-8. In addition, the mhb and hyb genes (Table S6) were also discovered in B-8,

452

indicating that lignin-derived salicylate and 3-hydroxybenzoate was degraded through

453

gentisate pathway. Moreover, the significant expression of the paa genes (Table S7)

454

also demonstrated the conversion of phenylacetate by the phenylacetyl-CoA

455

ring-cleavage pathway. The involvement of the phenol degradation pathway was also

456

noted due to the up-regulation of the phl genes (Table S8). The over-expressed mhp

52

presented in the lignin degradation by

22


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457

indicated the presence of the 2,3-dihydroxyphenylpropionate catabolic pathway

458

(Table S9).

459

All the samples exhibited the up-regulated lignin degradation genes as compared

460

to the control. The lignin depolymerization genes were significantly up-regulated in

461

the NaOH-treated RS samples, whereas the lignin-related aromatic compounds

462

degradation genes were significantly up-regulated in the APL samples. This

463

phenomenon is primarily due to the different lignin structures in APL and the RS

464

residue following NaOH pretreatment.

465

Currently, the degradation of lignin-related aromatic compounds not only

466

provides substrates for the tricarboxylic acid cycle but also acetyl-CoA for PHA

467

synthesis,3,

468

pathway) represent the most common pathways to supply precursors for PHA

469

synthesis.53 Herein, the transcriptome analysis revealed the presence of a type II

470

pathway 54 in B-8 due to the up-regulation of a gene cluster consisting of phaC, phaA,

471

phaB and phaR (Table S10). Moreover, the over-expressed fad genes indicated the

472

presence of the fatty acid β-oxidation pathway. The phaJ gene encoded enoyl-CoA

473

hydratase to channel hydroxyacyl-CoA from β-oxidation to PHA biosynthesis.55 A

474

gene similar to phaJ was up-regulated, thereby characterizing β-oxidation pathway as

475

another route for PHA precursor provision. The fatty acid de novo biosynthesis was

476

also involved on the up-regulation of the fab genes. 3-hydroxyacyl-ACP-CoA is

477

generally converted to 3-hydroxyacyl-CoA by 3-hydroxyacyl-ACP-CoA transacylase

17

while fatty acid de novo biosynthesis and metabolism (β-oxidation

23



Page 24 of 43

478

(PhaG), which is the enzyme that links fatty acid de novo biosynthesis and PHA

479

synthesis. However, the present study did not observe phaG gene neither from

480

transcriptome nor from the genome of B-8. Therefore, the fatty acid de novo

481

biosynthesis pathway may not be involved in the biosynthesis of PHA in B-8.

482

CONCLUSIONS

483

In summary, the present study proposed a novel platform of in-situ

484

bioconversion by B-8 to overcome the inherent challenges of delignification for

485

lignocellulose valorization. The biocatalysis promoted the digestibility of the RS to

486

achieve almost complete carbohydrate conversion. Meanwhile, the lignin fractions

487

were converted to PHA. The presented chemical and molecular analyses

488

demonstrated

489

depolymerization of lignin, and the subsequent metabolism of the released

490

lignin-related aromatic compounds, which then converted the intermediate acetyl

491

coenzyme A to PHA. This strategy provided a new perspective for lignocellulose

492

valorization.

493

ASSOCIATED CONTENT

494

Supporting Information

495

The Supporting Information is available free of charge on the ACS Publications

496

website.

that

B-8

secreted

extracellular

oxidative

24


enzymes

for

the

Page 25 of 43 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


497

PHA extraction methods; The location of genes and gene clusters,

498

carbohydrate utilization of B-8 (Figure S1-2); XRD, GPC, FTIR results of

499

the lignin fractions in RS and APL (Figure S3-7); Fluorescence imaging,

500

GC-MS, FTIR, H-NMR, GPC, TGA, and DSC results of the PHA (Figure

501

S8-13); The effect of the nitrogen limitation on the biocatalysis in

502

pretreatment (Figure S14); Heatmap of the transcriptional changes (Figure

503

S15); Comparison of biopretreatments with other research (Table S1);

504

Assignments of FTIR spectra for lignin (Table S2); Comparison of

505

production of PHA with other research (Table S3); The enzymes related to

506

lignin depolymerization (Table S4); Genes responsible for lignin-related

507

aromatic compounds pathways (Table S5-10)

508

Corresponding Author

509

Yan Shi

510

*E-mail:

511

+86-0731-88830875

512

Notes

513

The authors declare no competing financial interest

514

ACKNOWLEDGMENTS

[email protected];

Fax:

+86-0731-88710171;

25


Tel:


515

This work was supported by Program for Changjiang Scholars

516

(T2011116), National Funds for Distinguished Young Scientists of China

517

(50925417), National Natural Science Foundation of China (31400115).

26


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518

TABLES AND FIGURES

519

Table 1. Characterization of the lignin fractions in RS by GPC, 2D-NMR. Sample

RS

NaOH-RS

Co-tread RS

Mw (Daltons)

3105

1399

868

Mn (Daltons)

373

175

171

PDI

8.33

7.98

5.06

S/G ratio a

1.28

0.82

2.09

β-O-4 content b

86.3%

88.6%

100%

β-β content b

6.4%

-

-

β-5 content b

7.3%

11.4%

-

520

a

521

1

522

b

523

1

524

IA, IB, IC represent the α-position signal of β-O-4, β-β, and β-5, respectively;

525

Co-treated RS: NaOH+B-8 treated RS

S/G

ratio

=

Ix%=

0.5IS2,6/IG2

Ix/(IA+IB+IC)×100%

27



526 527

Figure 1. (a) Effect of the pretreatment conditions on the enzymatic hydrolysis of the

528

pretreated RS, Co-treated: NaOH+B-8 treated; (b) Effect of the pretreatment

529

conditions on the chemical composition of the pretreated RS, wherein the circular

530

images at the top represent the photos of the corresponding RS of each condition,

531

wherein the circular images at the top represent the photos of the corresponding RS of

532

each condition; (c) Relationship between the lignin content (%) and the sugar yield

533

(mg g-1) for a series of RS samples.

28


Page 28 of 43

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534 535

Figure 2. (a) SEM images of the different RS samples (a1: untreated RS, a2: NaOH-treated RS, and a3: Co-treated RS: NaOH+B-8 treated RS);

536

(b) Conceptual illustration of the proposed microstructure of RS, respectively; (c-e): AFM amplitude images and 3D images, where Ra is the

537

average surface area roughness; Rmax is the maximum vertical distance between the highest and lowest data points in the AFM image. 29



538 539

Figure 3. 2D-HSQC spectra of the lignin fractions obtained from the RS samples. (a)

540

Aromatic region of the untreated RS; (b) aromatic region of the NaOH-treated RS;

541

(c) aromatic region of the co-treated RS, co-treated RS: NaOH+B-8 treated RS; (d)

542

side-chain region of the untreated RS; (e) side-chain region of NaOH-treated RS; (f)

543

side-chain region of the co-treated RS; Main structures present in RS. (A) β-O-4

544

alkyl-aryl ethers; (B) resinols; (C) phenylcoumaran; (I) p-hydroxycinnamoyl alcohol

545

end groups; (FA) ferulates; (PCE) p-coumarates; (S) syringyl units; (S’) oxidized

546

syringyl units bearing a carbonyl at C; (G) guaiacyl units; (H) para-hydroxy-phenyl

547

units; (X) β-D-xylopyranoside.

30


Page 30 of 43

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548

549 550

Figure 4. 2D-HSQC spectra of the lignin fractions obtained from the APL. (a-b)

551

aromatic region of APL, (c-d) side-chain region of APL.

31



552 553

Figure 5. Removal of absorbance at 280 nm and total phenols from APL by B-8.

32


Page 32 of 43

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554 555

Figure 6. The elucidation of the lignin valorization mechanisms in B-8. The up-regulated genes were marked in red.

33



556

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For Table of Contents Use Only

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Synopsis

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In-situ bioconversion is first introduced to pretreatment strategy and this biocatalysis

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provides a new platform for lignocellulose valorization.

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43


In Situ Lignin Bioconversion Promotes Complete Carbohydrate

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