THESE DE DOCTORAT DE

L'UNIVERSITE DE NANTES

COMUE UNIVERSITE BRETAGNE LOIRE

ECOLE DOCTORALE N° 600 Ecole doctorale Ecologie, Géosciences, Agronomie et Alimentation Spécialité : « Biochimie, biologie moléculaire et cellulaire »

« Assessing the activity of a modular AA9 LPMO on cellulosic substrates » « Analysis of the soluble and the insoluble fractions» Thèse présentée et soutenue à Nantes, le 07 février 2020 Unité de recherche : BIA : Biopolymères, interactions, assemblages Thèse N° :

Par

Amani Chalak

Rapporteurs avant soutenance : Gabrièle Véronèse Directrice de recherche INRAE Toulouse Evangelos Topakas Associate Professor National technical University of Athens

Composition du Jury :

Président : Charles Tellier Professeur Université de Nantes Examinateurs : Gabrièle Véronèse Directrice de recherche INRAE Toulouse

Evangelos Topakas Associate professor National Technical University of Athens Caroline Rémond Professeur Université de Reims Champagne Ardenne

Dir. de thèse : Bernard Cathala Directeur de recherche INRAE Nantes Co-dir. de thèse : Jean-Guy Berrin Directeur de recherche INRAE Marseille Encadrante : Ana Villares Chargée de recherche INRAE Nantes

Thank you..

to Jean-guy, for your supervising all along the thesis. Thank you for your availability, trust,

encouragement, guidance and patience.

to Bernard Cathala and Ana Villares for your support, supervising and availability.

to Céline Moreau for guiding me in the QCM-D and NMR experiments.

to the Nano team for all the knowledge I have acquired.

to Craig Faulds for hosting me in the BBF laboratory and for all your encouragement and

support.

to Mireille Haon and Sacha Grisel for guiding me in the production and purification of the

enzymes and for the ionic chromatography measurements. Thank you for your availability,

and kindness.

To BBF lab members for the excellent work atmosphere, and for your kindness.

to the jury members Gabrièle Véronèse, Evangelos Topakas, Caroline Rémond, Charles

Tellier for accepting to evaluate this work.

to the thesis committee members Gabriel Paes and Julien Bras for following the progress of

my thesis work. Their comments and advice gave me the opportunity to see more globally the

orientation of my thesis and to be confident that it is going in the right direction.

to INRA and Region Pays de la Loire for financing this work.

To the platform BIBS for collaborating in this work

Thanks also to Angélina d’Orlando who helped me learn a lot in the Atomic Force

Microscopy analysis

to my friends Nancy Ramia and Mariane Daou for always being there to support and

encourage me especially when science didn’t reward me with good results.

“A friend is someone who understands your past, believes in your future, and accepts you just

the way you are.”

to my sister and my mother for your love and support, especially in the stressful moments of

the thesis.

“To us, family means putting your arms around each other and being there.”

Publications

Influence of the carbohydrate-binding module on the activity of a fungal AA9 lytic

polysaccharide monooxygenase on cellulosic substrates. Chalak A, Villares A, Moreau C,

Haon M, Grisel S, d’Orlando A, Herpoël-Gimbert I, Labourel A, Cathala B, and Berrin

J-G. Biotechnology for biofuels, 2019, 12(1):1-10.

Conference abstracts for posters

1-Third International EPNOE Junior Scientists Meeting:

Lytic polysaccharide monooxygenases (LPMOs) as novel tools for the

preparation of innovative nanocelluloses

Amani Chalak1 , Ana Villares

1 , Céline Moreau

1 , Mireille Haon

2 , Sacha Grisel

2 , Aurore

Labourel2, Jean-Guy Berrin

2 & Bernard Cathala

1

Cellulose is the most abundant biopolymer on Earth. It has been used for centuries for paper,

textile and chemicals production (Klemm et al., 2005). Nowadays, efficient breakdown of

cellulose is taking on a new importance as it constitutes a promising source for biofuels and

advanced bio-based products such as nanocelluloses. Nanocelluloses arise from the

fractionation of the cellulose fibers by chemical hydrolysis (cellulose nanocrystals) (Ranby,

1951) or mechanical delamination (cellulose nanofibers, NFC). In the case of NFC

production, to pass from the macroscale to the nanoscale, mechanical treatments require high

energy inputs that hinder efficient production of NFC. Lytic polysaccharide monooxygenases

(LPMOs) are recently discovered oxidative enzymes that belong to the AA9 family within the

CAZy database. When applied to cellulosic fibers, LPMOs facilitate the defibrillation down to

nanoscale, opening the road for a new pretreatment process (Villares et al,2017; Bennati

Granier et al., 2015). The objective of this study is to understand the LPMO action on

cellulosic fibers. For this goal, the LPMO9H from Podospora anserina was produced in

Pichia pastoris with and without its carbohydrate binding module (CBM), and the purpose of

this work is to investigate the contribution of the CBM domain in the activity of this LPMO.

In order to depict the level of hierarchy on which LPMOs act, two nanocellulosic materials

were used as substrates: Bacterial cellulose nanocrystals (BCNCs) as a crystalline cellulose

substrate, and regenerated cellulose as an amorphous cellulosic substrate. The action of

LPMO was investigated, in combination with cellobiohydrolase, by high performance anionic

exchange chromatography (HPAEC) and quartz crystal microbalance with dissipation (QCM-

D). Revealing the mechanism of action of LPMO is a key building block for their application

in the production of novel nanocellulosic materials in a way that respects the environment.

Bennati-Granier C., Garajova S., Champion C., Grisel S., Haon M., Zhou S., Fanuel

M., Ropartz D., Rogniaux H., Gimbert I., Record E., and Berrin J-G Biotechnology for

Biofuels 2015, 8:90

Klemm D., Heublein B., Fink H.P., and Bohn A. Angewandte Chemie International

Edition 2005, 44 : 3358 – 3393

Ranby B. G., Discuss. Faraday Society., 1951

Villares A., Moreau C., Bennati-Granier C., Garajova S., Foucat L., Falourd X., Saake

B., Berrin J.G. & Cathala B. Scientific Reports 2017, 7:40262 1BIA, INRA, 44300, Nantes, France.

2Biodiversité et Biotechnologie Fongiques, INRA,

Aix Marseille Univ, UMR1163, 13009, Marseille, France.

2-LPMO symposium

Influence of the CBM1 on the activity of the PaLPMO9H on cellulosic

substrates

A. Chalak

1,2, A. Villares

1, C. Moreau

1, M. Haon

2, S. Grisel

2, I. Gimbert

2, A. Labourel

2, B.

Cathala1, J.G. Berrin

2

1Biopolymères Interactions Assemblages, INRA, Nantes, France

2Biodiversité et Biotechnologie Fongiques, UMR1163, INRA, Aix Marseille Université,

Marseille, France

Cellulose-active lytic polysaccharide monooxygenases (LPMOs) secreted by filamentous

fungi play an important role in the degradation of recalcitrant lignocellulosic biomass. On a

biotechnological point of view, they could be used as innovative tools for the production of

nanocelluloses. However, their direct action on cellulosic substrates is not fully understood

(Villares et al., 2017). Model cellulosic substrates (nanofibrillated cellulose, amorphous and

crystalline celluloses) were used to probe the action of the LPMO9H from Podospora

anserina (PaLPMO9H ; Bennati-Granier et al., 2015) on cellulose. The carbohydrate binding

module (CBM1) of PaLPMO9H was deleted to assess its impact on the activity of the

enzyme. The deletion of the CBM1 impaired the release of soluble sugars and the binding to

cellulose. Synergy experiments with the cellobiohydrolase I (CBHI) from Trichoderma reesei

revealed that the truncated enzyme was still able to improve the efficiency of the CBHI in the

same range as the wild-type enzyme (PaLPMO9H). The cellulolytic action of the truncated

enzyme was further confirmed using complementary approaches (HPSEC, microscopy)

evaluating the action of the PaLPMO9H with and without CBM1 on the insoluble fraction of

cellulosic substrates. For this purpose, Kraft pulp fibers were used and the fibers’structural

modifications were evaluated by optical and atomic force microscopy. Preliminary analyses

indicate that the CBM1 does not preclude the activity of the enzyme on cellulose but its

presence has an important role in helping the enzyme to bind to the substrates and release

more soluble sugars. Understanding the level of hierarchy on which LPMOs act will be a

stepping stone in the design of novel nanocellulosic materials.

References

[1] A. Villares, C. Moreau, C. Bennati-Granier, S. Garajova, L. Foucat, X. Falourd, B. Saake,

J.G. Berrin and B. Cathala (2017). Scientific Reports 7:40262

[2] C. Bennati-Granier, S. Garajova, C. Champion, S. Grisel, M. Haon, S. Zhou, M. Fanuel,

D. Ropartz, H. Rogniaux, I. Gimbert, E. Record, and J.G. Berrin (2015). Biotechnology for

Biofuels, 8:90

3-CBM13 meeting

Influence of the carbohydrate binding module on the activity of a fungal

AA9 lytic polysaccharide monooxygenase

Amani Chalak1,2

, Ana Villares1, Céline Moreau

1, Mireille Haon

2, Sacha Grisel

2, Isabelle

Gimbert2, Aurore Labourel

2, Bernard Cathala

1,*, Jean-Guy Berrin

2,*

1Biopolymères Interactions Assemblages, INRA, Nantes, France

2Biodiversité et Biotechnologie Fongiques, UMR1163, INRA, Aix Marseille Université,

Marseille, France

E-mail: Amani.Chalak@inra.fr

Keywords: Podospora anserina, cellulosic substrates, LPMO9H, CBHI, Carbohydrate

Binding Module 1, CBM1, HPSEC, microscopy, Atomic force microscopy.

Background: Cellulose-active lytic polysaccharide monooxygenases (LPMOs) secreted by

filamentous fungi play an important role in the degradation of recalcitrant lignocellulosic

biomass (Bennati-Granier et al., 2015). They can occur as multidomain proteins fused to a

carbohydrate-binding module (CBM). On a biotechnological point of view, LPMOs are

promising and innovative tools for the production of nanocelluloses and biofuels (Villares et

al., 2017) but their direct action on cellulosic substrates is not fully understood.

Results: In this study, we probed the action of the family 1 CBM (CBM1) appended to the

LPMO9H from Podospora anserina (PaLPMO9H) using model cellulosic substrates. As

expected, the deletion of the CBM1 weakened the binding to nanofibrillated cellulose,

amorphous and crystalline cellulose. Although the release of soluble sugars from cellulose

was drastically reduced under standard conditions, the truncated LPMO retained some activity

on soluble oligosaccharides. The cellulolytic action of the truncated LPMO was demonstrated

using synergy experiments with a cellobiohydrolase (CBH). Indeed, the truncated LPMO was

still able to improve the efficiency of the CBH on cellulose nanofibrils in the same range as

the full length LPMO. Analysis of the insoluble fraction of cellulosic substrates evaluated by

optical and atomic force microscopy confirmed that the CBM1 module was not strictly

required to promote the disruption of the cellulose network. Based on these results we reduced

the amount of water in the reaction to increase the probability of enzyme-substrate interaction

in a CBM-free context. Increasing the substrate concentration enhanced the performance of

PaLPMO9H without CBM in terms of products release. Interestingly, removing the CBM

altered the regioselectivity of PaLPMO9H with a significant release of C1 oxidized products.

Conclusion: The absence of the CBM1 does not preclude the activity of the LPMO on

cellulose but its presence has an important role in driving the enzyme to the substrate and

releasing more soluble sugars (both oxidized and non-oxidized) therefore facilitating the

detection of LPMO activity at low substrate concentration. These results will help us to guide

the selection of suitable LPMOs for the production of nanocelluloses and biofuels.

________________

[1] Villares A., Moreau C., Bennati-Granier C., Garajova S., Foucat L., Falourd X., Saake B.,

Berrin J.G. and Cathala B. Scientific reports 2017 7, 40262

[2] Bennati-Granier C., Garajova S., Champion C., Grisel S., Haon M., Zhou S., Fanuel M.,

Ropartz D., Rogniaux H., Gimbert I., Record E., and Berrin J.G. Biotechnology for Biofuels,

2015 8, 90

Context ....................................................................................................................................... 1

I. Bibliographic introduction………………………………………………………………...3

I.1. The cellulosic biomass ..................................................................................................... 5

I.1.1. Introduction ............................................................................................................... 5

I.1.1.1. History and discovery of cellulose ......................................................................... 5

I.1.1.2. Origin of cellulose .................................................................................................. 5

I.1.2. Cellulose in plant cell wall ........................................................................................ 6

I.1.3. Plant cell wall composition ....................................................................................... 7

I.I.3.1. Molecular structure of cellulose ............................................................................. 8

I.1.3.2. Hemicelluloses, pectin and lignin .......................................................................... 8

I.1.4. Hierarchical organization of cellulose ...................................................................... 9

I.1.5. Cellulosic substrates ................................................................................................ 11

I.1.5.1. Phosphoric acid swollen cellulose (PASC) .......................................................... 11

I.1.5.2. Bacterial cellulose nanocrystals (BCNs) and cellulose nanocrystals (CNCs) ..... 12

I.1.5.3. Nanofibrillated cellulose (NFC) .......................................................................... 14

I.1.5.4. Kraft pulp fibers ................................................................................................... 18

I.2. Enzymatic degradation of cellulose ............................................................................... 19

I.2.1. General classification .............................................................................................. 19

I.2.2. Glycoside hydrolases .............................................................................................. 19

I.2.3. Oxidative enzymes .................................................................................................. 20

I.2.3.1. Discovery of LPMOs ........................................................................................... 21

I.2.3.2. Fungal enzymatic degradation of cellulose .......................................................... 21

I.2.3.3. Current classification of LPMOs ......................................................................... 22

I.II.3.4. Representative structures of LPMO families ...................................................... 25

I.2.3.5. Activity & mechanism of action of LPMO. ......................................................... 27

I.2.3.6. Mode of action of LPMOs on insoluble substrates .............................................. 28

I.2.3.7. Biotech applications using LPMOs ..................................................................... 30

References……………………………..………………………………………….............….33

II .Materials and methods…………………………………………………………………...41

II.1. Cellulosic substrates ..................................................................................................... 43

II.1.1. Phosphoric acid swollen cellulose (PASC) ........................................................... 43

II.1.2. Bacterial microcrystalline cellulose (BMCC) ....................................................... 43

II.1.3 Nanofibrillated cellulose (NFC) ............................................................................. 44

II.1.4. Delignified kraft paper ........................................................................................... 45

II.1.5. Cellooligosaccharides ............................................................................................ 45

II.2. Heterologous production of enzymes in Pichia pastoris .............................................. 45

II.2.1. Strains and media ................................................................................................... 45

II.2.2. Synthetic genes and vectors used .......................................................................... 45

II.2.3. Preparation of the competent cells of Pichia pastoris ........................................... 46

II.2.4. Transformation of Pichia pastoris ......................................................................... 47

II.2.5. Recombinant production of recombinant enzymes ............................................... 47

II.2.6. Enzyme purification .............................................................................................. 48

II.3. Protein analysis ............................................................................................................. 48

II.3.1. Dosage of protein ................................................................................................... 48

II.3.2. Protein electrophoresis .......................................................................................... 48

II.3.3. ICP-MS analysis .................................................................................................... 49

II.4. Amplex red assay ...................................................................................................... 49

II.5. Cellulose Binding Assays ............................................................................................. 49

II.6. Enzyme assays .............................................................................................................. 50

II.6.1. Combined assays ................................................................................................... 50

II.6.2. Analysis of oligosaccharides ................................................................................. 50

II.7. Analysis of the insoluble fraction ................................................................................. 51

II.7.1. Optical microscopy ................................................................................................ 51

II.7.2. Atomic force microscopy (AFM) .......................................................................... 51

II.7.3. Transmission electron microscopy (TEM). ........................................................... 51

II.7.4. NMR analysis of the NFC samples: Sample preparation ...................................... 52

II.7.5. Fourier transform infrared spectroscopy (FTIR) ................................................... 52

II.7.6. Neutral sugar composition ..................................................................................... 52

II.7.7. Colorimetric assay ................................................................................................. 53

II.7.8. High Performance Size exclusion Chromatography coupled with Multi Angle

Light Scattering detector (HPSEC-MALLS) ................................................................... 53

II.7.8.1. Enzymatic treatment ........................................................................................... 53

II.7.8.2. Fiber dissolution ................................................................................................. 54

II.7.8.3. Chromatographic analysis .................................................................................. 54

II.7.9. Quartz crystal microbalance with dissipation monitoring (QCM-D) .................... 54

References ............................................................................................................................... 56

Chapter III. Preparation and characterization of the substrates and the enzymes…….57

III.1 Characterization of substrates used in this study: ......................................................... 59

III.1.1. Bacterial Microcrystalline Cellulose (BMCC) ..................................................... 59

III.1.2.Nanofibrillated cellulose (NFC) ............................................................................ 61

III.1.2.1.Dry weight measurement: ................................................................................... 61

III.1.2.2.Optical microscopy: ............................................................................................ 62

III.1.2.3.Transmission electronic microscopy (TEM): ..................................................... 64

III.1.2.4. Carbohydrate composition: ................................................................................ 67

III.1.2.5.Solid state NMR: ................................................................................................. 69

III.1.2.6.Infrared analysis (FTIR): .................................................................................... 71

III-2 Recombinant production of the enzymes ..................................................................... 72

III.2.1 Introduction: ........................................................................................................... 72

III.2.1.1. Podospora anserina ............................................................................................ 72

III.2.1.2. Podospora anserina LPMO enzymes ................................................................. 73

III.2.1.3.The PaLPMO9H enzyme .................................................................................... 73

III.2.1.4.The roles of CBMs in cellulose-acting enzymes ................................................ 74

III.2.2. Results ................................................................................................................... 75

III.2.2.1. Heterologous production of PaLPMO9H with and without CBM1 .................. 75

Conclusion: ............................................................................................................................... 79

References…………………………………………………………………………………..…………80

Chapter IV. Depicting the activity of the LPMO enzyme by analyzing the soluble

fraction…………………………………………………………………………………… ….83

IV.1 Absence of CBM1 alters LPMO cellulolytic activity at low substrate concentration . 85

IV.2 LPMO-FL and LPMO-CD are both able to cleave cellohexaose ................................ 87

IV.3 The CBM1 favors the binding of LPMO to cellulosic substrates ................................ 88

IV.4 Combined action of LPMO-FL and LPMO-CD with a cellobiohydrolase .................. 89

IV.5 Increasing substrate concentration reduces the need for the CBM1 ............................ 90

References……………………………………………………………………………………93

Chapter V

Depicting the activity of the LPMO enzyme by analyzing the insoluble fraction ............ 95

V.1 Direct visualization of fiber morphology by microscopy ............................................. 97

V.2.Evaluation of chain length by high performance size exclusion chromatography

(HPSEC): ............................................................................................................................ 100

V.2.1.The weight average molecular weight of the non-mechanically-treated samples 100

V.2.2.The weight average molecular weight of the mechanically-treated samples ....... 101

V.2.3.The number average molecular weight and the polydispersity of the non-

mechanically-treated samples ........................................................................................ 102

V.2.4.The number average molecular weight and the polydispersity of the mechanically-

treated samples ............................................................................................................... 103

V.3. Real-time monitoring of action of LPMO on regenerated cellulose by quartz crystal

microbalance with dissipation monitoring (QCM-D) ........................................................ 104

V.3.1.The QCM-D technique ......................................................................................... 105

V.3.2.Preparation of the regenerated amorphous cellulose layer ................................... 106

V.3.3.Control experiment: Injecting cysteine or ascorbate and CBHI to the regenerated

amorphous cellulose surface .......................................................................................... 106

V.3.4.Injecting LPMO enzymes and CBHI enzyme to the regenerated amorphous

cellulose surface ............................................................................................................. 109

V.4 Detecting the oxidative cleavage at the surface of cellulose using a colorimetric method

............................................................................................................................................ 110

V.4.1. Assessment of the method on Avicel and PASC ................................................ 110

V.4.2. Testing of LPMO-FL and LPMO-CD enzymes on PASC .................................. 111

References ............................................................................................................................. 113

Chapter VI. Discussion…………………………………………………………………….115

VI.1. Importance of substrates to study LPMOs ............................................................ 117

VI.2. Importance of experimental methods to assay LPMO activity ............................. 118

VI.3. Importance of experimental conditions to assay LPMO activity .......................... 119

VI.4. Depicting the impact of the CBM for LPMO action............................................. 119

VI.1. Which LPMOs for nanocelluloses production? .................................................... 120

References…………………………………………………………………………………..122

Annexe……………………………………………………………………………...……….123

Résumé de la thèse en français ……………………………………………………………125

List of figures

Figure 1 Anselme Payen. ........................................................................................................... 5

Figure 2 Three-dimensional structure of the secondary cell wall of a tracheid. ........................ 7

Figure 3 Molecular structure of cellulose................................................................................... 8

Figure 4 Schematic representation of crystalline and amorphous regions of cellulose

microfibrils. .............................................................................................................................. 10

Figure 5 From cellulose sources to the cellulose molecules.. .................................................. 11

Figure 6 TEM images of dried dispersion of cellulose nanocrystals. ...................................... 13

Figure 7 Mechanical processes for NFC production. ............................................................... 15

Figure 8 (a) Schematic illustration of CNFs and CNCs production from fiber cell walls by

mechanical and chemical treatments, respectively .................................................................. 17

Figure 9 Optical microscopy image of kraft fibers .................................................................. 18

Figure 10 Current view of the fungal enzymatic degradation of cellulose.. ............................ 22

Figure 11 Representative 3D structure of LPMO families ...................................................... 25

Figure 12 Structural views of the Lentinus similis AA9 LPMO with the copper ion depicted as

a sphere.. ................................................................................................................................... 26

Figure 13 Structural views of the principal protein contacts between cellotriose and the

binding surface of LsAA9 in LsAA9-G3 structure. ................................................................. 26

Figure 14 LPMOs catalyze oxidation leading to chain cleavage……………………………..28

Figure 15 Solid state 13C CP/MAS NMR spectra of bleached softwood kraft pulp. .............. 29

Figure 16 Aspect of NFC obtained by microfluidization after LPMO pretreatment ……… .. 31

Figure 17 Set-up of the production of BMCC………………………………………………..44

Figure 18 Description of the expression vector pPICZαA……………………………….. ….46

Figure 19 TEM images of stained bacterial microcrystalline cellulose. ............................................... 60

Figure 20 Infrared spectrum of BMCC substrate .................................................................................. 60

Figure 21 Optical microscopy image of Ex Piano (2%) at different scales ........................................... 62

Figure 22 Optical microscopy image of Ex Piano (10%) at different scales......................................... 63

Figure 23 Optical microscopy image of NFC (2.5%) CTP at different scales. ..................................... 64

Figure 24 TEM microscopy of ex piano 2%. ........................................................................................ 65

Figure 25 TEM microscopy of ex piano 10% ....................................................................................... 66

Figure 26 TEM microscopy of nanofibrillated cellulose 2% (NFC CTP)............................................. 67

Figure 27 NMR full spectra of the three cellulosic substrates .............................................................. 69

Figure 28 NMR C1 peak deconvolution of the three cellulosic substrates. .......................................... 70

Figure 29 NMR C4 peak deconvolution of the three cellulosic substrates. .......................................... 71

Figure 30 Infrared spectra of the four samples of nanocelluloses ......................................................... 72

Figure 31 Schematic representation of the enzymes used in this study. ............................................... 75

Figure 32 Production of LPMO-CD in two different types of competent cells: ................................... 76

Figure 33 Comparison of the sizes of the three different truncated LPMOs. ........................................ 77

Figure 34 Fluorimetric assay for the generation of hydrogen peroxide.........................................……79

Figure 35 Analysis of soluble degradation products. ............................................................................ 86

Figure 36 Time -course analysis and quantification of the Glc4 and Glc3. .......................................... 87

Figure 37 Qualitative cellulose binding assays. .................................................................................... 89

Figure 38 Combined action of LPMO-FL and LPMO-CD with a cellobiohydrolase (CBH). .............. 90

Figure 39 Analysis of degradation products generated by LPMO-FL and LPMO-CD. ....................... 91

Figure 40 Morphology of LPMO-treated kraft fibers. ............................................................. 99

Figure 41 Mw average of the non-mechanically-treated samples ........................................... 101

Figure 42 Mw average of the mechanically-treated samples .................................................. 102

Figure 43 Number-average molecular weight and polydispersity index of set #1 and set#2 of

the non-mechanically-treated samples. .................................................................................. 103

Figure 44 Number-average molecular weight and polydispersity index of set #1 and set#2 of

the mechanically-treated samples. .......................................................................................... 104

Figure 45 Schematic representation of the decreasing of the signal for the QCM-D

measurements on a rigid layer adsorbed on the surface with low dissipation, and a flexible and

viscoelastic layer with high dissipation. ................................................................................. 106

Figure 46 The frequency and dissipation monitoring of the QCM-D regenerated cellulose

surface. ................................................................................................................................... 107

Figure 47 The frequency and dissipation monitoring of the QCM-D regenerated amorphous

cellulose surface. .................................................................................................................... 108

Figure 48 The frequency and dissipation monitoring of the QCM-D regenerated amorphous

cellulose surface where LPMO-FL was injected. .................................................................. 109

Figure 49 Absorbance of the (PV)-Ni2+

complex present in the supernatant of different

samples treated or not with PaLPMO9E. ............................................................................... 111

Figure 50 Absorbance of (PV)-Ni2+

present in the supernatant of different samples with

LPMO-FL and LPMO-CD. .................................................................................................... 112

List of tables

Table 1: Summary of the principal characteristics of the existing LPMO families….……….24

Table 2: Average dry weight measurement of the three different types of NFC samples…...61

Table 3: weight % of sugars in the three samples of nanocelluloses…………………………68

Abbreviations

3D: three dimensional

AA: Auxiliary activity

AFM: Atomic force microscopy

AGU: Anhydroglucose unit

AOX1: alcohol oxidase 1 gene

AS: Accessible surface

BMCC: bacterial microcrystalline cellulose

BMGY: Buffered complex medium containing glycerol

BMMY: Buffered Methanol-complex Medium

C1: Carbon 1

C4: Carbon 4

C4-ox: oxidized at C4

CAZy: Carbohydrate-active enzyme database

CBH: cellobiohydrolase

CBM: Carbohydrate binding module

CDH : Cellobiose dehydrogenase

CE : Carbohydrate esterases

CNC: Cellulose nanocrystals

CNF: cellulose nanofibers

Cr: Crystalline form

CTP : Centre technique du papier

DMAc: Dimethylacetamide

DNA: Deoxyribonucleic acid

DP: Degree of Polymerization

DTT: Dithiothreitol

EC: enzyme commission

EG: endoglucanase

FAD: Flavin adenine dinucleotide

FTIR: Fourier Transform Infrared Spectroscopy

GH: glycoside hydrolases

Glc 2: cellobiose

Glc 3: cellotriose

Glc 4: cellotetraose

Glc 5: cellopentaose

Glc 6: cellohexaose

Glc2ox: oxidized cellobiose

GPC: gel permeation chromatography

HEPES: 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

Hj AA9 A: Hyprocrea jecorina auxiliary activity 9-C

HPAEC/PAD: High-performance Anion-exchange chromatography coupled with pulsed

amperometric detection

HPSEC-MALLS: High-performance size-exclusion chromatography coupled with on-line

multi-angle laser light scattering

IAS : Inaccessible surface

ICP-MS: Inductively Coupled Plasma Mass Spectrometry

IgG-like fold: Immunoglobulin-like domains

IUBMB: International union of biochemistry and molecular biology

L :Length

LFAD: Lateral fibril aggregate dimension

LFD: Lateral fibril dimension

LPMO: Lytic Polysaccharide Monooxygenases

LPMO-CD: Lytic Polysaccharide Monooxygenase-catalytic domain

LPMO-FL: Lytic polysaccharide Monooxygenase-full length

m/z: mass/charge

Mw: molecular weight

Mw/Mn: polydispersity

Nc AA9-C: Neurospora crassa auxiliary activity 9-C

NcAA9-F: Neurospora crassa auxiliary activity 9-F

NFC: Nanofibrillated Cellulose

NMR Nuclear magnetic resonance

His: Histidine

OD optical density

P. anserina: Podospora anserina

PAD: Pulsed Amperometric detection

PaLPMO9E: Podospora anserina lytic polysaccharide monooxygenase 9E

PaLPMO9H: Podospora anserina lytic polysaccharide monooxygenase 9H

PASC: Phosphoric Acid Swollen Cellulose

PCr: Paracrystalline

Phe: Phenylalanine

PMMA 60: Polymethyl methacrylate 60

PMMA 95: Polymethyl methacrylate 60

PTFE membrane: Polytetrafluoroethylene membrane

PV: Pyrocatechol Violet

QCM-D: Quartz Crystal Microbalance with Dissipation monitoring

S layer: Secondary layer

SDS-PAGE: sodium dodecyl sulfate–polyacrylamide gel electrophoresis

solid state C13 CP/MAS/NMR: 13C Solid-state nuclear magnetic resonance spectroscopy

TEM: transmission electron microscopy

TEMPO: 2,2,6,6-tetramethylpiperidine-1-oxyl

Tris-HCl: hydroxymethyl aminomethane hydrochloride

Tyr: Tyrosine

v/w: volume/weight

w/v: weight/volume

YPD medium: yeast extract peptone dextrose medium

1

Context

In the new era of our world, we have increasing scarcity of oil resources and we depend on

petroleum-based polymers, which cause serious environmental pollution. Based on these

facts, non-toxic natural polymeric materials have attracted considerable attention from

reseachers, corporations, and governments. Cellulosic materials and nanocellulosic materials

derived from lignocellulosic biomass have been studied thoroughly over the last few years.

They are renewable, eco-friendly and unfold unique and interesting properties that make these

materials prone to be used at large scale in various applications. Nowadays, in the frame of

the bioeconomy, many countries are shifting their economies in a way that is respectful to

nature and are using less chemical agents. On a biotechnological point of view, using nature’s

microorganisms in biotechnological processes or getting inspired by the processes they adapt

to their natural habitats is a common trend. Using the microorganisms in biotechnological

processes or trying to use their processes leads to the reduction of the chemicals use and their

resulting toxic effects.

Chapter I

Bibliographic introduction

5

I.1. The cellulosic biomass

I.1.1. Introduction

I.1.1.1. History and discovery of cellulose

Cellulose is the most abundant, natural, renewable, and biodegradable polymer on Earth. It

represents about 1.5×1012

tons of the total annual biomass production. As such, it could satisfy

the increasing demand for environmentally-friendly and biocompatible products (Klemm et

al. 2002); (Kaplan 1998); (Klemm et al. 2005). Cellulose was discovered and isolated by the

French agricultural chemist Anselme Payen in 1838 (Figure 1). He went to study woods and

during his research, he began to separate out a substance resembling starch. He named this

substance, which he found in abundance in the cell walls of all plants he studied, cellulose

(Payen (1838); (Payen 1842); (Phillips 1940).

Cellulose was used, in history, for several purposes production of rope and cordage,

papermaking, writing documents, and making propellants and gun powder (Hon 1994).

I.1.1.2. Origin of cellulose

Cellulose is widely distributed in higher plants, in marine animals (for example tunicates) and

in non-plant sources including algae, fungi, bacteria, invertebrates and even amoeba

(protozoa) (Klemm et al. 2005). Other plants also contain a large amount of cellulose,

including hemp, flax, jute, ramie and cotton (Eichhorn et al. 2010). By far, the most

commercially exploited natural resource containing cellulose is wood. Cellulose can be

isolated from other agricultural residues as corn cob, corn stalks, rice straw, sugarcane

bagasse and wheat straw by using several techniques such as steam explosion, treatment with

sodium hydroxide, treatment with hydrogen peroxide or treatment with sulfuric acid or with

Figure 1 Anselme Payen.

6

acetic acid. The pulps obtained have degrees of polymerization (DPs) and crystallinity index

that vary according to their sources or the treatment used (Heinze et al. 2018).

I.1.2. Cellulose in plant cell wall

Cellulose is one of the most important primary plant component, yet its biosynthesis pathway

is not fully understood (Brown et al. 1996). The plant cell wall consists of a rigid layer of

complex polysaccharides on the outer surface of the plasma membrane that encases the plant

cell (Dey and Brinson 1984);(Fry 1988), and it is one of the most important characteristic

features of plant cells. It is the main system for osmoregulation and provides the strength and

rigidity to the cell and the plant, allowing growth and expansion of the cell (Brett and

Waldron 1996); (Dey and Brinson 1984); (Reiter 2002). The mature cell wall is a complex

laminate structure of three distinct layers, the middle lamella, the primary cell wall and

secondary cell wall.

Primary cell walls continue to be deposited throughout the growing stage of the cell. When

the developing cell reaches its definitive size, the secondary wall is formed between the

primary wall and the cell membrane. It is composed of three different layers, S1, S2, and S3

(Figure 2), and it is important in providing mechanical strength for the tissue (Timell 1986).

The cellulose microfibrils of the S2 and S3 layers are aligned in an ordered, parallel

arrangement, which differs from S2 layer to S3 layer. These layers contain also

hemicelluloses and lignin. The three S layers can be modified during cell maturation and the

amount of lignin and cellulose laid down in the secondary wall is influenced by biological

origin or abiotic factors such as mechanical stress (Plomion et al. 2001).

7

Figure 2 Three-dimensional structure of the secondary cell wall of a tracheid (xylem

cells).

The composition of cell walls changes between the different cell types. The development or

the environmental conditions can change the make-up of cell-walls between different cell

types (Wang et al. 2016).

I.1.3. Plant cell wall composition

The primary cell wall is mainly composed of cellulose (15-30%), hemicelluloses (25%;

xyloglucan, xylan and glucomannan), pectins (30%; homogalacturonan, rhamnogalacturonan

type I, rhamnogalacturonan type II), proteins (20%), and phenolic acids (ferulic, coumaric).

Another “primary wall”is formed when the secondary wall start its formation. It is the

transformation of the primary cell wall in a layer included in the secondary wall. This

“primary wall” of the secondary wall is highly lignified and there is almost no pectin. The

cellulose microfibrils are randomly positioned and linked to hemicelluloses by hydrogen

bonds that help the positioning of the microfibrils. In addition to this complex structure, the

arrangement of the cellulose microfibrils is not homogeneous across the thickness of the

primary wall. The secondary wall is mostly composed of cellulose (50-85%), hemicelluloses

(0-25%) and lignin (0-25%) depending on plant species. Components are linked together by

hydrogen bonds (cellulose-hemicelluloses) and crosslinked in the case of lignin-cellulose or

lignin-hemicelluloses. The cellulose microfibrils are arranged helically as discussed in the

preceding chapter on cell wall structure (Plomion et al. 2001).

8

I.I.3.1. Molecular structure of cellulose

Since its discovery by Payen, the physical and chemical aspects of cellulose have been

intensively studied. Cellulose is a carbohydrate homopolymer formed of repeating units of β-

D-glucopyranose molecules that are covalently linked through glycosidic bonds between the

equatorial OH group of the C4 and the C1 carbon atom (β-1,4-glucan) (Figure 3).

The chemical repeating unit is therefore the D-glucopyranose molecule and the structural

repeating unit is cellobiose, formed by two adjacent monosaccharides (DP2). To

accommodate the preferred bond angles of the acetal oxygen bridges, every second

anhydroglucose unit (AGU) ring is rotated 180° in the plane. Each AGU has three hydroxyl

groups. As a result, cellulose has a large number of hydroxyl groups on its extensive linear

chain forming the polymer. These groups and their ability to form strong hydrogen bonds

confer to cellulose its most important properties: (i) multi-scale microfibrillated structure, (ii)

hierarchical organization, and (iii) highly cohesive nature (Lavoine et al. 2012); (Klemm et al.

2005).

I.1.3.2. Hemicelluloses, pectin and lignin

Hemicelluloses represent a large group of heteropolymers displaying β-(14) linked

backbones. According to the different cell walls, hemicelluloses can be divided into four

general classes of polysaccharide types: xyloglucans, xylans, mannans and glucomannans

and β-glucans with mixed linkages. These types of hemicelluloses can be found in the cell

wall of terrestrial plants except for β-(13, 14) glucans which are restricted to Poales and a

few other groups. The detailed structure of hemicelluloses varies widely between species and

cell types. The most important biological role of hemicelluloses is their contribution to

strengthening the cell wall by interacting with cellulose and in some walls with lignin.

Hemicelluloses are synthesized by glycosyl transferases located in the Golgi membranes.

(Scheller and Ulvskov 2010).

Figure 3 Molecular structure of cellulose (n=DP, degree of polymerization).

9

Another constituent of plant cell wall is pectin. Pectins are heteropolysaccharides comprising

homogalacturonan, rhamnogalacturonan I and II. They are abundant in walls that surround

growing and dividing cells, walls of cells in the soft parts of the plant, and in the middle

lamella and cell corners. Pectin is also present in the junction zone between cells and

secondary walls including xylem and fiber cells in woody tissue (O'Neill et al. 2004).

Lignin is the generic term for a large group of aromatic polymers resulting from the oxidative

combinatorial coupling of 4-hydroxyphenylpropanoids (Boerjan et al. 2003); (Ralph et al.

2004). These polymers are deposited predominantly in the walls of secondarily thickened

cells, making them rigid and impervious. In addition to developmentally programmed

deposition of lignin, its biosynthesis can also be induced upon various biotic and abiotic stress

conditions, such as wounding, pathogen infection, metabolic stress, and perturbations in cell

wall structure (Caño‐Delgado et al. 2003); (Tronchet et al. 2010); (Vanholme et al. 2010).

I.1.4. Hierarchical organization of cellulose

The cellulose chain is formed by a certain number of AGU. The DP, which is the number of

AGU per chain, varies upon the cellulose source and treatments. For cotton, DP can be higher

than 10,000. Depending on the severity of cooking and pre-treatment, cellulose used in wood

pulps has an average DP of 600 to 1200 and 100 to 200 for cellulose powders, such as

microcrystalline cellulose prepared by acidic hydrolysis. Cellulose chains are polydisperse

and the molecular mass distribution strongly depends on sources.

The three hydroxyls groups (OH groups) of the glucose unit can form hydrogen bonds by

intra and inter molecular interactions. The strength of these hydrogen bonds is around 25

kJ/mol, which are responsible of the stiffness of the cellulose molecule. Intramolecular

hydrogen bonds are formed between O-3-H and O-5’ and between O-2-H and O-6’of adjacent

glucose units. Intermolecular hydrogen bonds are formed between O-6-H and O-2’ and

between O-6-H and O-3’. Intramolecular hydrogen bonds are partly responsible for the linear

integrity and rigidity of the polymer chain and intermolecular hydrogen bonds result in

crystalline structures and other supramolecular arrangements. The elementary fibril or

microfibril contains several cellulose chains bound by intermolecular hydrogen bonds. Within

the microfibril all the (OH) groups are equatorial, with the methane (CH) groups oriented to

the ring axially, which results in the appearance of hydrophilic regions parallel to the ring

plane and hydrophobic regions perperdicular to the ring (Habibi et al. 2010). Elementary

fibrils are packed into larger units called microfibrillated cellulose. The microfibrillated

10

celluloses are assembled into cellulose fibers which are represented in Figure 5. The

elementary fibril has a diameter of about 5 nm and several micrometers in length whereas the

microfibrillated cellulose has a diameter ranging from 20 to 50 nm and a length of several

µm. Each microfibril is formed with cellulose crystals linked by disordered amorphous

regions (Azizi Samir et al. 2005). The ordered regions are stabilized by a strong and complex

network of hydrogen bonds (Habibi et al. 2010) (Figure 4). The disruption of the highly

packed structure results in the release of nanocelluloses. There are two main types of

nanocelluloses: (i) cellulose nanocrystals and (ii) cellulose microfibrils (Lavoine et al. 2012).

Figure 4 Schematic representation of crystalline and amorphous regions

of cellulose microfibrils (Rajinipriya et al. 2018).

11

Figure 5 From cellulose sources to the cellulose molecules: details of the cellulosic fiber

structure with emphasis on the cellulose microfibrils (Lavoine et al. 2012).

I.1.5. Cellulosic substrates

I.1.5.1. Phosphoric acid swollen cellulose (PASC)

PASC is a homogeneous fraction of cellulose obtained by swelling cellulose in

orthophosphoric acid and blended to homogeneity. The procedure was firstly developed by

Walseth (Walseth 1952) to produce highly reactive cellulose suitable for cellulase activity

studies. Since then, PASC has become one of the most common cellulose substrates for

enzymatic assays. Phosphoric acid pretreatment has been used to create cellulose samples of

various surface areas. The acid treatment opens larger holes on the surface of plant cell walls

by removing the easily digested lignocellulose fraction (e.g., hemicellulose) and separating

the fibers. Depending on the cellulose origin and the concentration and reaction time,

phosphoric acid can swell or dissolve cellulose. Hence, recent studies using phosphoric acid

to increase cellulose accessibility in lignocellulosic samples suggested the presence of a

critical point in the phosphoric acid concentration above which cellulose was completely

dissolved (80-85%, depending on the cellulose origin) (Moxley et al. 2008); (Zhang et al.

2006). Swelling in phosphoric acid disrupts orderly hydrogen bonds in crystalline cellulose

resulting in an amorphous arrangement of chains.

12

I.1.5.2. Bacterial cellulose nanocrystals (BCNs) and cellulose nanocrystals

(CNCs)

Bacterial cellulose is obtained from species belonging to the genera of Acetobacter,

Rhizobium, Agrobacterium, and Sarcina through various cultivation methods and techniques,

the most studied being Acetobacter xylinum. The reason why the bacteria generate cellulose is

unclear, but it has been suggested that it is necessary for their survival, such as to guard

against ultraviolet light, or to act as a barrier to fungi, yeasts and other organisms. Unlike

plant cellulose, bacterial cellulose is synthesized in its pure form without lignin, hemicellulose

and pectin. It is produced in the form of gel-like, never dry sheets containing up to 99% of

water. The resulting microfibrils are microns in length and have a large aspect ratio (defined

as the length-to-width, L/w) greater than 50 with a morphology depending on the specific

bacterial and culturing conditions. Its structure is composed of crystalline and amorphous

zones. The latter can be hydrolyzed by acid treatment to isolate BCNs.

Cellulose nanocrystals (CNCs) were firstly produced by Bengt G. Rånby (Rånby 1951) by

treating bleached and refined sulfite pulp with sulfuric acid. When appropriate combination of

acid concentration, time and temperature are used, the amorphous regions of the cellulose

fiber are hydrolyzed faster than crystalline ones allowing the isolation of CNCs. (Habibi et al.

2010). Sulfuric acid hydrolysis is the most common technique because it produces cellulose

crystals that are dispersible in water due to the sulfate ester groups introduced on the surface

during hydrolysis. Other acids, such as hydrochloric acid or phosphoric acid have also been

investigated for preparing CNCs (Camarero Espinosa et al. 2013). CNCs are rigid, rod-like

cellulose nanoparticles of high aspect ratio whose dimensions are around 5-20 nm in width

(w) and 50-2000 nm in length (L), depending on their biological origin. Hence, cotton CNCs

are 5-10 nm in width and nanocrystals from tunicates present width around 5-30 nm and

length of several microns (Lin et al. 2012); (Moreau et al. 2016); (Sacui et al. 2014) (Figure 6)

13

Figure 6 TEM images of dried dispersion of cellulose nanocrystals derived from (a)

tunicates, (b) bacteria, (c) ramie and (d) sisal (Habibi et al., 2010).

Bacterial cellulose nanocrystals (BCNs) present a high aspect ratio with rectangular cross

sections or a ribbonlike shape with dimensions of 10 nm × 50 nm and a range in length from

100 nm to several micrometers.(Araki and Kuga 2001) (Pirich et al. 2015) It is possible to

hydrolyze BCN at mild conditions and avoid the use of sulfuric acid which promotes highly

charged surfaces. The use of hydrochloric acid results in charge-free nanocrystals (Pirich et al.

2015) so that BCNs appear to be bundled or aggregated, which can be due to the lack

electrostatic repulsion.

Typically, bacterial cellulose presents crystallinity values around 63% (Moon et al. 2011).

The removal of amorphous regions by acid hydrolysis increases crystallinity to 70-95%, (Park

et al. 2010) which is high compared to other sources of CNCs, such as cotton or wood (54-

88%) (Moon et al. 2011).

BCNs have shown unique properties such as mechanical strength, barrier properties, high

water retention to be exploited as reinforcing agents in nanocomposites, edible and

biodegradable food packaging, membranes, or gas barriers. Furthermore, they have

demonstrated excellent compatibility for applications as scaffold for tissue engineering,

antimicrobial films, tablet excipients, and drug delivery systems. (Reiniati et al. 2017). In

addition, BCNs act as emulsion stabilizers in a wide variety of applications including

paintings, cosmetics, and food industry, among others(Capron et al. 2017; Kalashnikova et al.

2011).

14

I.1.5.3. Nanofibrillated cellulose (NFC)

NFC was firstly produced by Turbak in 1983 by mechanical delamination of cellulose fibers

(Turbak et al. 1983). Fibrillation releases long flexible and semi-crystalline NFC consisting of

alternating crystalline and amorphous regions, which are around 5-20 nm in width and several

microns in length. NFC presents a high aspect ratio and forms strongly entangled and

disordered networks and gels. Even if a substantial part of the non-crystalline domains remain

essentially intact, the degree of crystallinity of NFC is usually high.

Presently, there are a large number of different mechanical disintegration processes to

produce NFC. They may be sub-divided into conventional (homogenization, microfluidization

and refining) and non-conventional process (extrusion, steam explosion, ball milling

ultrasonification, aqueous counter collision etc.). The most commonly used is the high

pressure homogenization, which consist of passing the cellulose slurry through a tiny gap

between the homogenizing valve and an impact ring (Figure 7), subjecting the fibers to shear

and impact forces. Likewise, the principle of the microfluidizer consists in passing the

cellulose suspension through a thin chamber with a specific geometry with an orifice width of

100–400 µm. By applying high pressure, strong shear forces and impact of the suspension

against the channel walls are achieved. Other strategies, such as ultra-fine grinding process

where the cellulose slurry is passed between static and rotating grinding stones (disks) are

also used. (Nechyporchuk et al. 2016).

15

Figure 7 Mechanical processes for NFC production. Adapted from www.niro-soavi.com,

www.microfluidicscorp.com and www.masuko.com (Nechyporchuk et al. 2015)

The major impediments in the NFC production are the high energy consumption, and the

clogging of the homogenizer. The energy consumption during disintegration is related to the

cohesion of the cell wall; and the clogging phenomena are related to the aggregation of the

fibers. Both of these factors are affected by the colloidal interactions (Lindstrom 2017).

In order to reduce the energy involved in mechanical delamination of cellulose fibers,

different pre-treatments have been developed, namely enzymatic and chemical treatment. The

production of NFC became a combination of different operations that included the mechanical

treatment as the last step (Paakko et al. 2007).

- Enzymatic pretreatment: It was demonstrated that a combination of high pressure shear

forces and mild enzymatic pretreatment constitutes an efficient method to prepare NFC with a

well controlled diameter in the nanometer range and to maintain high aspect ratio, in contrast

to acid hydrolysis. Enzymatic pre-treatments involve the use of endoglucanases, which

degrade the fiber and facilitate their delamination; therefore, reducing the need for mechanical

treatment (Bombeck et al. 2016).

16

- Chemical treatment: The decrease of energy consumption of the mechanical treatment can

be done by the introduction of charged groups into the fiber walls. The most used route is the

catalytic oxidation using 2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO). This treatment

consists in the oxidation of the secondary hydroxy groups at C6 OH present on each fibril

surface to carboxylate groups using NaBr and NaClO as an additional catalyst and primary

oxidant, respectively. (Saito et al. 2009) Repulsions between the charged groups favor the

fibrillation (Klemm et al. 2011). (Saito et al. 2006) demonstrated that the use of TEMPO-

mediated oxidation on never-dried cellulose resulted in the disintegration of the fibers into

individual microfibrils by a simple mechanical treatment. They proved that a combination of

TEMPO-mediated oxidation and mechanical treatment can produce microfibrils with smaller

widths, using a much lower energy input (Saito et al. 2006). Another chemical pre-treatment

is the carboxymethylation, which is the introduction of carboxymethyl groups on the cellulose

surface (Walecka 1987). The charge repulsion leads to a drastic decrease in fiber–fiber

friction and therefore the fibers have less susceptibility to flocculate and less clogging

tendency (Horvath and Lindström 2007). Nevertheless, the TEMPO oxidation usually

involves side reactions that depolymerize the cellulose chain, and high quantities of toxic

wastes.

NFC exhibits high stiffness (138 GPa), low thermal expansion (0.1 ppm/K) and low density

(1.6 g/cm3) (Saito et al. 2009). Its exceptionally large specific surface area implies potential

increased interactivity with secondary components/materials. This could create innovative

breakthroughs in use of these materials in areas such as e.g. nanocomposites, packagings,

coatings and dispersion technology (Stenstad et al. 2008) Furthermore, NFC shows

extraordinary optical, mechanical, and thermal properties. (Wei et al. 2014). These features

contribute to their applications such as reinforcement components in flexible display panels

and oxygen-barrier layers. The synergistic use of such nanofibers together with proper

functionalization pathways can produce materials with diverse properties namely aerogels,

emulsions, templated materials and stimuli-responsive nanodevices (Tingaut et al. 2012).

Also, these materials can be strong carriers for the incorporation of guest nanomaterials. The

resulting nanocellulose-based nanocomposites show many advantages including antimicrobial

and catalytic activities that can be applied for the purification of water. Moreover,

nanocellulose can be a substrate for fuel cell fabrication and, thus, shows promise in energy

applications (Wei et al. 2014).

17

Figure 8 (a) Schematic illustration of CNFs and CNCs production from fiber cell walls

by mechanical and chemical treatments, respectively (Salas et al. 2014). (b) Schematics

of idealized cellulose fibers showing one of the suggested configurations of the crystalline

and amorphous regions, and CNCs after sulfuric acid hydrolysis of the amorphous

regions, exhibiting the characteristic sulfate half ester surface groups formed as a side

reaction (Domingues et al. 2014). (c) Systematic diagram of preparation of CNFs by

surface carboxylation using TEMPO oxidation (Chirayil et al. 2014). (Chaker and Boufi

2015) (Du et al. 2019)

18

I.1.5.4. Kraft pulp fibers

In the paper industry, the fabrication of pulp consists of separating the cellulose fibers from

the other cell wall components with the lowest impact for the fiber. This separation can be

achieved by mechanical treatments or by chemical reactions, as well as by a combination of

both. The kraft pulping is the most common form of chemical pulping, at 80% of the total

chemical pulping industry. It is an alkaline process that comprises the treatment of wood with

a hot mixture of water, sodium hydroxide, and sodium sulfide, which is known as white

liquor, at temperatures between 170°C and 175°C. The treatment breaks the bonds between

lignin, hemicellulose and cellulose. The baking time is between two and five hours according

to the nature of wood. This step is used to separate the fibers one from the other by dissolving

lignin. For a chemical pulp, the middle lamellae is eliminated by dissolution of the lignins and

because of this, the fibers are only slightly shortened unlike mechanical pulp where fibers

undergo an important physical treatment that shortens them strongly. This process gives the

most resistant unbleached pulp, with a yield of around 50% (Chevalier-Billosta 2008).

Kraft pulp fibers (Figure 9) have a width of 15-50 μm and several milimeters in length.

During kraft pulping, lignin is dissolved whereas variable amounts of hemicelluloses remain,

depending on the biomass source.

Figure 9 Optical microscopy image of kraft fibers

19

I.2. Enzymatic degradation of cellulose

Cellulose-acting enzymes are secreted in nature by many microorganisms, mainly bacteria

and filamentous fungi. The most widely used microorganism in industry is the fungus

Trichoderma reesei that secretes high level of a cellulase blend able to degrade efficiently

cellulose. Several fungi other than T. reesei (e.g., Myceliophthora thermophila, Aspergillus

niger) have also been reported to be excellent cellulase producers (Gusakov 2011).

I.2.1. General classification

Carbohydrate-active enzymes were initially classified according to their activity and substrate

specificity in the classification of the International Union of Biochemistry and Molecular

Biology (IUBMB) with attribution of corresponding Enzyme Commission (EC) numbers. In

the 1990’s, cellulose-acting enzymes have been classified within the carbohydrate-active

enzyme (CAZy; www.cazy.org) database (Lombard et al. 2014). This sequence-based

classification underpins all functional, structural and mechanistic consideration of

carbohydrate-active enzymes, which are grouped by families within the glycoside hydrolases

(GH) that cleave glycosidic bonds, the carbohydrate esterases (CE) that allow de-acylation of

sugars and the Auxiliary Activity enzymes (AA) that gathers mainly oxido-reductases.

Carbohydrates Binding Modules (CBM) are non-enzymatic modules which are associated to

CAZymes to promote their binding. Enzymes belonging to the same CAZy family may have

different substrate specificities and reciprocally several enzymes acting on the same substrate

can be classified in different families. In October 2019, the CAZy database included 165 GH

families, 16 CE families, 16 AA families and 85 CBM families.

I.2.2. Glycoside hydrolases

Glycoside Hydrolases (GHs) (EC 3.2.1.-) are a widespread group of enzymes, which

hydrolyse the glycosidic bond between two or more carbohydrates. The hydrolysis of the

glycosidic bond is catalyzed by two amino acid residues of the enzyme: a general acid (proton

donor) and a nucleophile/base. Enzymatic hydrolysis of cellulose requires the combined

action of at least three types of GHs (endoglucanases (EC 3.2.1.4), cellobiohydrolases (EC

3.2.1.176) and beta-glucosidases (EC 3.2.1.21)), capable to hydrolyse the β-1,4 covalent

bonds that connect glucose units in the cellulose fiber. These cellulases act synergistically on

cellulose chains with different specificities.

20

Endoglucanases (EG; endo-1,4 β-D-glucanases) perform the random endo-hydrolysis of

(1→4)-β-D-glucosidic linkages in cellulose, lichenin and cereal β-D-glucans. Enzymes with

EG activity are encountered in several CAZy families (GH5, GH6, GH7, GH9, GH12, GH45,

GH74 and GH131) and display various structures such as (β/α)8 barrels, jelly rolls or 7-fold β-

propellers.

Exoglucanases (cellulose 1,4-β-cellobiosidases, EC 3.2.1.91), also known as

cellobiohydrolases (CBH) perform the hydrolysis of (1→4)-β-D-glucosidic linkages in

cellulose and similar substrates, in a processive manner releasing cellobiose from the reducing

or non reducing ends of the chains. Enzymes with CBH activity are mainly encountered in

CAZy families GH6 and GH7.

Beta-glucosidases (1,4-β-glucosidase) perform the hydrolysis of cellobiose or longer β-D-

glucosyl oligosaccharides released upon action of the two other types of cellulases and release

β-D-glucose residues. Enzymes with beta-glucosidase activity are mainly encountered in

CAZy families GH1 and GH3.

Different characteristics of the cellulose influence the efficiency of enzymatic hydrolysis such

as the degree of polymerization (DP), crystallinity, particle size and surface area. Regarding

the crystallinity for example, it was shown that it has a strong impact on the rate of hydrolysis.

In general, it is considered that the enzymatic attack preferentially occurs in amorphous

regions, which should lead to an increase of the degree of crystallinity in remaining cellulose,

but several studies have shown that this index is not strongly affected during hydrolysis

(Penttila et al. 2010).

The synergistic action of the different types of cellulases is an essential phenomenon in the

hydrolysis of cellulose. Two types of synergies have been identified, one endo-exo between

endoglucanases and cellobiohydrolases (Jalak et al. 2012) and one exo-exo between two

cellobiohydrolases one acting on reducing end and the other one acting on non-reducing end

(Teeri 1997).

I.2.3. Oxidative enzymes

In addition to the classical cellulose-acting glycoside hydrolases described above, it is now

recognized that other enzymes play a role in the degradation of cellulose. Oxidative enzymes

acting on polysaccharides have been recently added to the CAZy database in the AA class

21

(Levasseur et al. 2013) following the discovery of lytic polysaccharide monooxygenases

(LPMO), which needed a reclassification of these families into a suitable category.

I.2.3.1. Discovery of LPMOs

The two families that formed LPMOs originally, in CAZy, were the GH family 61 (GH61)

comprising fungal enzymes originally proposed to have weak endoglucanase activity

(Morgenstern et al. 2014); and the CBM family 33 (CBM33) including mainly bacterial

proteins binding chitin. A functional link was found between the two families by comparative

structural analyses of bacterial CMB33 (Vaaje-Kolstad et al. 2005) and fungal GH61

(Karkehabadi et al. 2008) members. GH61s and CBM33 were evidenced to be oxidative

enzymes (Vaaje-Kolstad et al. 2010) using copper as a redox catalytic metal (reviewed by

(Hemsworth et al. 2015)).

Addition of GH61 and gallate or ascorbate significantly increased the hydrolysis of cellulose

by cellulases (Quinlan et al. 2011); (Beeson et al. 2012); (Westereng et al. 2011). In 2012,

they were described as lytic oxidases and then the term ‘lytic polysaccharide monooxygenase’

(LPMO) was collectivelly adopted to describe them (Horn et al. 2012). The word ‘lytic’

illustrates the ability of these enzymes to break and loosen polysaccharide chains.

When CAZy reclassified redox carbohydrate active enzymes into auxiliary activity families

(AA), GH61s and CBM33s were reassigned as families AA9 and AA10, respectively.

I.2.3.2. Fungal enzymatic degradation of cellulose

Most cellulolytic microorganisms secrete a large set of enzymes when grown on cellulosic

substrates, among which are the enzymes with auxiliary activities (mostly LPMOs) that act

synergistically with cellulases (see figure 10).

22

Figure 10 Current view of the fungal enzymatic degradation of cellulose. Note that many

cellulolytic enzyme systems have multiple EG and/or CBH that may act on various parts

of the substrate, e.g. different crystal faces or parts differing in terms of crystallinity and

accessibility. Cellobiose dehydrogenase (CDH) may provide AA9 LPMOs with electrons,

but it must be noted that not all organisms have genes encoding for both of these enzyme

families in their genome. Also other non-enzymatic reductants (electron donors) have

been demonstrated to induce oxidative activity (e.g. reduced glutathione, ascorbic acid,

gallic acid, lignin) (Horn et al. 2012).

I.2.3.3. Current classification of LPMOs

Since their discovery in 2010, new LPMO families have emerged. Most LPMOs have been

identified using a “module walking” approach. This method relies on the fact that many

CAZymes are multi-modular with one or more additional domains, which are often substrate-

targeting CBMs. The modules attached to known CAZymes were used to search for proteins,

which (i) contained those modules, (ii) contained a conserved histidine immediately after the

signal peptide cleavage site and (iii) displayed insignificant sequence similarity to known

AA9 and AA10 families. This method has led to the discovery of several LPMO families

(AA11, AA13, AA15) but did not allow to identify new LPMOs lacking additional modules.

Using comparative post-genomic analyses, AA14 and AA16 were discovered and

characterized (Couturier et al. 2018; Filiatrault-Chastel et al. 2019).

Today, LPMOs (recently assigned EC 1.14.99.53-56) are grouped in seven CAZy families

(AA9-AA11 and AA13-AA16), which are differentiated by their amino acid sequences. These

families have been found in bacteria, viruses, fungi and recently in arthropod species (AA15

LPMOs) and in plants (see Table 1).

23

Fungal AA9 LPMOs are active on cellulose, cello-oligosaccharides and

hemicelluloses containing glucose units linked in β-1,4: xyloglucans, glucomannans, mixed β-

glucans (Isaksen et al. 2014) ; (Bennati-Granier et al. 2015);(Frommhagen et al. 2016)

(Fanuel et al. 2017) An AA9 LPMO from Myceliophtora thermophila has been shown to be

active on xylan, but only in the presence of cellulose (Frommhagen et al. 2015) and another

AA9 LPMO from Lentinus similis is able to cleave both xylan and xylo-oligosaccharides

(Simmons et al. 2017).

LPMOs from the family AA10, are mainly present in bacteria and few viruses and

they are active on chitin, cellulose and sometimes on both (Forsberg et al. 2014) (Agostoni et

al. 2017). Recently an AA10 from the actinobacterium Kitasatospora papulose has shown

activity on cellulose, chitin and xylan (Ribeiro Correa et al. 2019).

The families AA11 and AA13 were discovered in fungi due to sequence homologies

with domains carried by known LPMO families (Vu et al. 2014); (Hemsworth et al. 2014; Lo

Leggio et al. 2015). They both contain only a few characterized members at the moment, and

are respectively active on chitin and on starch constituents (amylose and amylopectin).

The AA14 LPMOs were discovered due to the development of secretomic approaches

(Berrin et al. 2017b). The characterization of two of them from Pycnoporus coccineus,

revealed that they are active on the xylan covering the fibers of cellulose in wood (Couturier

et al. 2018).

The family AA15 gather LPMO members active on cellulose and on chitin, from

various organisms: oomycetes, unicellular and multicellular algae, virus, terrestrial and

marine invertebrates (crustaceans, mollusks, insects, millipedes, spiders) (Sabbadin et al.

2018).

The catalytic modules of these different families are frequently associated with other modules

like the carbohydrate binding module (CBM) whose specificities correspond to substrates

identified for each family (see Table 1) (Lenfant et al. 2017); (Horn et al. 2012); (Berrin et al.

2017a) (Couturier et al. 2018; Vu et al. 2019) (Sabbadin et al. 2018).

24

Table 1: Summary of the principal characteristics of the existing LPMO families. The

informations are extracted from the CAZy database website and also from the different

references in the text.

CAZy

famil

y

Type of

organisms

Associated

CBM

Number of

characterize

d enzymes

Specificity of

substrate

Regioselectivit

y

AA9 Fungi

CBM1

(cellulose),

CBM18

(chitin)

37

Cellulose, cello-

oligosaccharides,

mixed glucans,

xyloglucan,

glucomanan,

xylan

C1, C4,

& C1/C4

AA10

Bacteria,

virus,

archaea,

plants, fungi

(Ustilaginaceae

)

CBM1

(cellulose),

CBM2

(cellulose

/chitin/xylan),

CBM12,

CBM5

(cellulose

/chitin),

CBM14

(chitin),

CBM20

(starch)

23 Cellulose, chitin,

xylan

C1 & C1/C4

(cellulose)

C1 (chitin)

AA11 Fungi None 2 Chitin C1

AA13 Fungi CBM20

(starch) 3 Starch C1

AA14

Fungi

(Pycnoporus

coccineus)

None 2 Xylan associated

to cellulose C1

AA15

Invertebrates

, oomycetes,

algae, virus

CBM1

(cellulose),

CBM14

(chitin)

2 Cellulose, chitin C1

AA16

Fungi

(Aspergillus

aculeatus)

CBM1

(cellulose) 1 Cellulose C1

25

Fungi are the organisms where we find the highest number of different LPMO families. They

have several genes of each family especially from the family AA9 where the number can

reach more than 30 (e.g. Podospora anserina). The reasons for this expansion are not well

known, but they could be linked to the diversification of the family in terms of substrate

specificity and other biochemical properties, and reflect differences in regulation that helps in

the adaptation to different substrates (Berrin et al. 2017a). The secretomic studies seem to go

in the same direction, since they demonstrate that only some of the LPMOs encoded by the

fungal genomes are secreted, and that they differ according to the substrate used to grow the

fungus (Berrin et al. 2017a). Amongst the enzymes secreted simultaneously, complementary

activities and regioselectivities are found for an efficient degradation of polysaccharides.

I.II.3.4. Representative structures of LPMO families

Figure 11 Representative 3D structure of LPMO families.

Each LPMO family AA9–AA11 and AA13–15 exhibit the recognizable Histidine brace

(Figure 11, top row, stick representation), composed of an N-terminal His (methylated in

fungal LPMOs) and an internal His for the equatorial Cu coordination. The His-brace is

completed with a Tyr residue in the axial position, sometimes replaced with a Phe residue in

some AA10 LPMOs. LPMO structures from all families have been solved with a Cu atom

(orange spheres) in the active site, with the exception of the newly discovered AA14 family

where a copper site similar to AA9 LPMOs was though confirmed by EPR spectroscopy.

LPMOs exhibit a common IgG-like fold (Figure 11, bottom row, cartoon representation) with

variation in surface topology (grey) depending on loop regions and helices. The catalytic Cu-

site is located at the surface of the substrate binding region (Tandrup et al. 2018).

26

A 3D structure of an AA9 LPMO in complex with its substrate was obtained by (Frandsen et

al. 2016). They reported the crystal structure of the Lentinus similis AA9 LPMO (LsAA9) in

complex with a cello-oligosaccharide (Figures 12 and 13). This LPMO’s structure reveals the

essential molecular features of LPMO interaction with substrates. For instance, the structures

revealed polar residues interacting with cellooligosaccharides near the active site and

demonstrated for the first time that a conserved Tyrosine located on the same surface was

involved in substrate binding. This information is essential to get better understanding of the

mechanism of action of these enzymes.

Figure 12 Structural views of the Lentinus similis AA9 LPMO with the copper ion

depicted as a sphere. a, Ribbon view of LsAA9; b. View of the histidine brace

coordinating the copper. (Frandsen et al. 2016).

Figure 13 Structural views of the principal protein contacts between cellotriose and the

binding surface of LsAA9 in LsAA9-G3 structure (Frandsen et al. 2016).

27

I.2.3.5. Activity & mechanism of action of LPMO.

The catalytic mechanism of LPMOs has been the centre of scientific debate. Monooxygenase

activity was first proposed in 2010 (Vaaje-Kolstad et al. 2010) and the mononuclear copper

active site was shown in 2011 (Quinlan et al. 2011). Likely events are hydrogen abstraction at

C1 or C4 of the substrate, followed by hydroxylation through an oxygen rebound mechanism

and elimination to the final product. Polysaccharide oxidation is accompanied by reduction of

molecular oxygen to water. Hydrogen abstraction from polysaccharides requires formation of

a very powerful oxidative species. In LPMOs the activation of oxygen is accomplished by

reduction of Cu(II) to Cu(I) and relocation of the electron to dioxygen upon binding to Cu(I)-

LPMO (Kjaergaard et al. 2014). Two electrons are needed in each cycle, one to reduce the

copper, and another for reduction of oxygen to water.

The electrons are provided by exogenous donors that can be naturally-occuring in

lignocelluloses (gallic acid, lignin), added externally (ascorbic acid, glutathion) or a co-

secreted enzyme such as CDH, a secreted flavocytochrome only occurring in fungi (Zamocky

et al. 2006). CDHs are composed of an N-terminal heme domain, which carries a cytochrome

b type heme and a C-terminal flavin domain which contains FAD, connected by a flexible

linker CDH is found in the genome of most wood-degrading fungi (Harreither et al. 2011).

Some of them also contain a C-terminal CBM1 targetting cellulose (Subramaniam et al.

1999). They catalyze the reducing end oxidation of cellobiose, cellodextrins or other

oligosaccharides to the corresponding lactones that are subsequently converted to their

aldonic acids. CDHs can act synergistically with LPMOs in cellulose hydrolysis (Wilson

2012);(Dimarogona et al. 2012) (Bey et al. 2013).

A futile side-reaction especially in the absence of natural substrate cellulose is the reduction

of O2 to H2O2 by LPMO. It has not been reported so far and is not the proposed in vivo

function. By adding different amounts of LPMO to the assay mixture they could prove that

the formation of H2O2 is directly proportional to the LPMO concentration. The formation of

H2O2 depends on the availability of a suitable reductant for the LPMO type-2 copper center

(Kittl et al. 2012).

28

I.2.3.6. Mode of action of LPMOs on insoluble substrates

As mentionned earlier, members of the different LPMO families display different substrate

specificities. AA9 have been extensively studied on cellulose by measuring the release of

soluble oligosaccharides. It allowed to differenciate differences in term of regioselectivity

either at the C1 or C4 or C1 and C4 carbon of the glucose (Hemsworth et al. 2014); (Bennati-

Granier et al. 2015). (Figure 14) But a striking difference between LPMOs and glycoside

hydrolases is the ability of LPMOs to cleave cellulose without the need to extract a cellulose

chain. Indeed, LPMOs are able to bind and directly cleave cellulose at the surface of fibers.

Figure 14 LPMOs catalyze oxidation within a polysaccharide chain leading to chain

cleavage Type-1 (C1) LPMOs oxidize at the C1 positon resuting in the formation of a

lactone, which is hydrated to generate an aldonic acid at the reducing end. Type-2 (C4)

LPMOs oxidize at the C4 position, resulting in the formation of a ketoaldose at the non-

reducing end (Adapted from (Hemsworth et al. 2015)).

Villares et al. investigations have been one of the first to show that an AA9 LPMO was able

to disrupt the cellulose fibers structure (Villares et al. 2017). NMR analysis of cellulose

treated with an AA9 LPMO (PaLPMO9H) revealed that chain breakage and associated

chemical modifications might induce weakening of the hydrogen bonds and van der Waals

network. This modification facilitates the mechanical delamination and/or creates new entry

H2O2

29

points for the attack by other hydrolytic enzymes, such as endoglucanases.The creation of

nicking points mostly at the non-crystalline part of the fibers weakens the cohesion of the

overall architecture of the fibers (Figure 15). Therefore LPMOs can be viewed as a novel

strategy for the production of nanofibers with high DP and crystallinity, and susceptible to be

further transformed into novel materials.(Villares et al. 2017).

Figure 15 Solid state 13C CP/MAS NMR spectra of bleached softwood kraft pulp. Leſt:

Deconvolution of C-4 region with crystalline forms Cr (Iα), Cr (Iβ) and Cr (Iα +β)

(black), para-crystalline form (PCr) (grey), accessible fibril surfaces (AS) (green), and

inaccessible fibril surface (IAS) (dark green) signals for reference cellulose fibers (a)

before and (b) aſter dispersion, and for cellulose fibers submitted to the LPMO

treatment (c) before and (d) aſter dispersion. Four signals from hemicelluloses and/or

cellulose oligomers are indicated in orange. Yellow line in (a,b,c and d) corresponds to

the sum of individual peaks resulting from the spectral deconvolution. In spectrum (d),

two additional peaks are denoted by (*) and (**) (see text for details). Right: Schematic

representation of cellulose fibril model for (e) reference cellulose fibers, and LPMO-

30

treated cellulose fibers (f) before and (g) aſter dispersion. The number of points in the

scheme is proportional of the area of the corresponding NMR signals. Blue arrows

represent possible nicking points created by the LPMO on the cellulose. Fibril scheme

displays the different cellulose forms: lateral fibril dimension (LFD), and lateral fibril

aggregate dimensions (LFAD).

In another LPMO single-molecules studies (Eibinger et al. 2014), it has been shown by

microscopic techniques (atomic force microscopy, AFM) that a NcAA9_F is localized to

crystalline regions of cellulose and thereby allows hydrolases to digest recalcitrant patches on

the substrate. A more recent study by the same group (Eibinger et al. 2017) shows co-

localization of NcAA9_C and NcAA9_F (one with C1 and one with C4 preference) on the

crystalline surfaces also targeted by a synergistic cellobiohydrolase. In contrast with a recent

AFM study of HjAA9_A on bacterial cellulose nanocrystals (Song et al. 2018), where the

LPMO showed a stop-and-go behavior, NcAA9_C and NcAA9_F appeared immobile. Using

synchrotron UV (ultraviolet) fluorescence imaging (Chabbert et al. 2017), it was shown that

LPMOs act in synergy with cellulases to degrade miscanthus plant cell walls, and synchrotron

infrared spectroscopy showed tissue-dependent effects. In terms of biotechnological

applications, in line with Villares et al. (2017), endoglucanase, LPMO and xylanases were

shown to facilitate nanofibrillation of paper pulp (Hu et al. 2018), potentially reducing the

need for mechanical refining while resulting in a pulp with a more uniform nanofibril

composition.

I.2.3.7. Biotech applications using LPMOs

The use of LPMOs in the production of biofuels :

The energy demand has considerably increased since the last century and problems are

associated with global warming including the rising of atmospheric greenhouse gas levels and

scarcity of fossil fuels. Therefore, it is imperative to reduce our heavy dependancy on fossil

fuels. Countries throughout the world search for new fuel alternative fuels including biofuels.

They have lower geenhouse gas emission, renewability and sustainability. Commercially,

there are many sources of biofuels such as second generation biofuels produced from

lignocellulosic biomass.It should be noted that lignocellulosic biofuels production suffers

from high production costs and other technical barriers due to the recalcitrance of biomass to

enzymatic degradation.

31

Given their flat surfaces, LPMOs are able to access the crystalline protion of polysaccharides

releasing new ends that can elicit the activity of canonical enzymes, promoting a boosting of

sugar release (Horn et al. 2012; Ribeiro Correa et al. 2019). LPMOs are now part of industrial

enzymatic cocktail, which are used for the production of second generation biofuels (Johansen

2016).

The use of LPMOs to defibrillate the cellulose fibers and to produce cellulose nanofibrils:

LPMOs are commonly used in synergy with cellulases to enhance biomass deconstruction.

However, there are few examples of the use of monocomponent LPMOs as a tool for

cellulose defibrillation. In the work of (Moreau et al. 2019), an AA9 LPMO was used as a

tool to produce nanofibrillated cellulose (NFC) based on the fact that LPMO action facilitates

the disruption of wood cellulose fibers. The fungal PaLPMO9E was used since it displays

high specificity toward cellulose and its recombinant production in bioreactor is easily

upscalable. Firstly, the treatment of birchwood fibers with PaLPMO9E resulted in the release

of a mixture of C1-oxidized oligosaccharides without any appartent modification in fiber

morphology and dimensions. The subsequent mechanical shearing disintegrated the LPMO-

pretreated samples yielding nanoscale cellulose elements. Their gel-like aspect and

nanometric dimensions demonstrated that LPMOs disrupt the cellulose structure and facilitate

the production of NFC (Figure 16) (Moreau et al. 2019).

Figure 16 Aspect of NFC obtained by microfluidization after LPMO pretreatment

(Moreau et al. 2019).

In another study (Valls et al. 2019), cellulases released cellooligosaccharides, reducing fiber

length and partially degraded cellulose. They also yielded 18% of nanofibrillated cellulose

(NFC) since they improved mechanical fibrillation. LPMOs introduced a slight amount of

COOH groups in cellulose fibers, releasing cellobionic acid to the effluents. LPMO improved

the action of cellulases. However, the COOH groups created disappeard from fibers. After

32

mechanical fibrillation of LPMO-cellulase treated cotton linters a 23% yield of NFC was

obtained. The combined treatment with LPMO and cellulose provided films with higher

transparency (86%), crystallinity (92%), smoothness and improved barrier properties to air

and water than films casted from non-treated linters and from commercial NFC.

In a follow up study from the same group (Valenzuela et al. 2019), cellulose nanofibrils were

produced using the AA10 LPMO from Streptomyces ambofaciens. The activity of the

bacterial LPMO showed high variability depending on the origin and degree of crystallinity of

the substrate. Additionally, they tested the effectiveness of SamLPMO10C in the

nanofibrillation of flax, a high crystalline agricultural fiber, as a single pretreatment or in

combination with cellulases. All pretreatments were followed by a mechanical defibrillation

by high-pressure homogenization to obtain cellulose nanofibrils. LPMO and cellulases

combined together showed higher fibrillation yield, optical transmittance eand carboxylate

content than control reactions. Therefore, LPMO could be explored as a promising green

alternative to reduce the energy consumption in the production of NFC (Valenzuela et al.

2019).

33

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Chapter II

Materials and methods

43

II.1. Cellulosic substrates

Several cellulosic substrates were used in this study. They represent either the crystalline,

amorphous, alternating crystalline and amorphous regions, or really close to natural fibers like

the kraft fibers.

II.1.1. Phosphoric acid swollen cellulose (PASC)

Phosphoric acid swollen cellulose (PASC) was chosen as the amorphous cellulosic substrate.

PASC is prepared by the swelling of cellulose in phosphoric acid so that cellulose chains are

separated and become disordered and more accessible. The protocol for producing PASC was

based on the method developed by Wood 1988 (Wood 1988) and described in the article of

Bennati-Granier et al., 2015 (Bennati-Granier et al. 2015). 10 g of Avicel (Fluka, Sigma-

Aldrich, Saint Louis, USA) were added to 200 mL of orthophosphoric acid (H3PO4, 85%

m/v), and the mixture was stirred at 4°C overnight. As cellulose swelling proceeded, viscosity

increased; therefore, water (800 mL) was added, and the mixture was stirred and

homogenized. Then, PASC was filtered through a Buchner filter vacuum filtration set-up and

successively washed by adding progressively a total of 5 L of milliQ water. Then,

approximately 400 mL PASC were neutralized by adding 500 mL NaHCO3 at 1% (pH 8.6)

four consecutive times and purified by vacuum filtration until obtaining a pH of neutralized

PASC of 7.The concentration of the PASC was later determined by the dry weight.

II.1.2. Bacterial microcrystalline cellulose (BMCC)

Bacterial microcrystalline cellulose (BMCC) was used as the crystalline cellulosic substrate.

Bacterial cellulose is a highly pure and crystalline material produced by several types of

aerobic bacterial species. It consists of highly ordered cellulose microfibrils arranged in a 3D

web-shaped structure. Compared to plant cellulose, bacterial cellulose has considerably higher

crystallinity (80-90%). BMCC was obtained from nata de coco cubes. Nata de coco is a gelly-

like food produced in Philippines. Its main concept consists in the fermentation of coconut

water through the production of microbial cellulose by Acetobacter xylinum. The cellulose is

composed of crystalline and amorphous zones. The latter can be hydrolyzed by acid treatment

so that only crystalline regions remain. The hydrolysis of the cellulose by hydrochloric acid

treatment allows obtaining a colloidal suspension composed of crystals of nanometric

dimensions. They form unstable colloidal suspensions since the hydrochloric acid does not

introduce charges on the surface of the nanocrystals.

44

Nata de coco cubes were firstly purified by dialysis to remove free sugars. After eight

dialyses, ultra-violet visible spectrophotometry was used to detect the presence of sugars or

impurities. Later, the cubes were mixed with waring blender to obtain a slurry. They were

filtrated to remove water using Buchner filtration 20CHR Whatman. We weighed the mass of

the solution to calculate the dry weight before acid hydrolysis. The experimental set up is

shown in Figure 17.

They were subjected to hydrochloric acid (2.5 N) hydrolysis at a temperature of 72°C in three

consecutive steps of total time of two hours, separated by filtration and three centrifugations

at 10,000 g for 10 minutes. Then, dialysis was done against distilled water.

Figure 17 Set-up of the production of BMCC.

II.1.3 Nanofibrillated cellulose (NFC)

Nanofibrillated cellulose (NFC) is obtained by mechanical delamination of cellulose fibers

which results in long flexible nanofibers consisting of alternating crystalline and amorphous

regions. Its diameter is in the range 10-60 nm and it has a length of several micrometers. The

NFC used in this thesis was kindly provided by the Centre Technique du Papier (CTP,

45

Grenoble, France). It was obtained via an endoglucanase pretreatment followed by

microfluidization.

II.1.4. Delignified kraft paper

Delignified bleached kraft pulp from resinous trees was used as the cellulose substrate. It

consists of fibers of several microns in diameter and some mm in length, containing both

crystalline and amorphous regions. Cellulose fibers were dispersed in 50 mM of sodium

acetate buffer and stirred for 48 h prior to enzymatic assays (Villares et al. 2017)

II.1.5. Cellooligosaccharides

The cellooligosaccharides used were purchased from Megazyme Wicklow, Ireland. They

were diluted in milliQ water to the concentration of 10 mM, aliquoted and stored at -20°C

until use.

II.2. Heterologous production of enzymes in Pichia pastoris

II.2.1. Strains and media

P. pastoris strain X33 and the pPICZαA vector are components of the P. pastoris Easy select

expression system (Invitrogen, Carslbad, USA). The SuperMan 5 strain of Pichia pastoris

produces recombinant proteins with less glycosylation (Pekarsky et al. 2018). In this

GlycoSwitch strain, glycan processing enzyme (OCH1) has been mutated to prevent glycan

elongation and a heterologous mannosidase is overexpressed to cleave extra glycans to

generate homogenous Man5 structure. All standard media and protocols are described in the

Pichia expression manual (Invitrogen). All media were autoclaved for 30 min at 110°C.

II.2.2. Synthetic genes and vectors used

The genes encoding PaLPMO9H was codon optimized for the expression in Pichia pastoris.

The genes (full length and truncated versions) were synthetized by Genewiz (South Plainfield

USA) and inserted into the vector pPICZαA (Figure 18) using BstBI and XbaI restriction

sites. The pPICZαA vector contains (i) the AOXI promoter inducible by the methanol, (ii) the

α-factor signal peptide allowing secretion of the recombinant enzyme, (iii) a Histidine tag at

the C-terminal position for the purification of the recombinant enzyme, (iv) a zeocin resistant

gene for the selection step, and (v) the sequences 5’ and 3’ of the gene aox1 for the integration

by homologous recombination in the genomic DNA of the yeast.

46

Recombinant expression plasmids were sequenced to check the integrity of the corresponding

sequences.

Figure 18 Description of the expression vector pPICZαA

II.2.3. Preparation of the competent cells of Pichia pastoris

The preculture of P. pastoris cells was prepared in 5 mL of YPD medium (10 g.L-1

yeast

extract; 20 g.L-1

peptone and 20 g.L-1

glucose) in a 50-mL Falcon tube. The tube was closed

with a slightly opened lid. The preculture was inoculated with cells from a petri dish. The

time of inoculation was 5-6 hours at 30°C and 160 rpm. The preculture was then diluted with

YPD medium to obtain an OD=0.45 at 600 nm. Two non-baffled flasks of 1 L containing 250

ml of YPD medium were inoculated by 2 mL of the dilution. The culture was incubated

overnight at 30°C and 200 rpm. The culture was stopped when the OD ranged between 1.3

and 1.5. The 2 x 250 mL volumes were centrifuged for 5 minutes at 4°C, 2000 g in sterile

centrifugation tubes. The cells were then resuspended in 100 ml of YPD medium buffered

with 20 ml of HEPES 1 M pH 8 and supplemented with 2.5ml of DTT 1M prepared

extemporaneously. The solutions were agitated very slowly and incubated at 30°C for 15

minutes and the volume was adjusted to 400 mL with cold sterile water. Then cells were

centrifuged at 4°C and 4000 g for 5 min and resuspended in 250 mL of cold sterile water.

Centrifugation was done again at 4°C at 4000g for 5 min and the cells were resuspended in 20

ml of cold sorbitol 1M. After a final centrifugation under the same conditions, cells were

resuspended in 500 µL of cold 1M sorbitol and aliquoted by fractions of 60 µL in small tubes

and stored at -80°C until use.

47

II.2.4. Transformation of Pichia pastoris

Pichia competent cells were thawed on ice. 0.5-5 µg of the linearized plasmid was placed at

the edge of the 2-mm gap electroporation cuvette (Eppendorf, Hambourg, Allemagne), and 60

µL of competent cells were added to the drop of DNA. The solution was allowed to migrate to

the bottom of the cuvette by tapping gently on the borders. The cuvette was placed on ice for

5 min. A 1500 V voltage was applied during 5 ms using a microspulsor electroporator (Biorad

Marned-la-Coquette, France). 1 mL of sorbitol (1 M) was added in the cuvette under sterile

conditions (near the Bunsen burner) at room temperature. Solutions were transferred into 15

mL tubes and incubated during two hours at 30°C without agitation. We prepared the Petri

dish of yeast extract peptone dextrose (YPD) medium (containing 10 g.L-1

yeast extract, 20

g.L-1

peptone, 20 g.L-1

dextrose and 20 g.L-1

agar) and add zeocine (500 µg mL-1

and 100 µg

mL-1

) to select the strains that have incorporated the plasmid. 100 µL of the electroporated

cells were spread on the petri dishes. The petri dishes were incubated at 30°C during 48 hours.

The transformants were then stored at -80°C in a medium containing 20% of sterile glycerol.

Zeocin-resistant P. pastoris transformants were then screened for protein production in

deepwell plates and recombinant enzymes were purified using an automated method.

Basically, 2 mL of YPD medium were inoculated in a 24-deepwell plate format by the

colonies from the petridish. The deepwells were incubated at 30°C under agitation at 200 rpm

and after 5-6 hours, 5 mL of BMGY medium was added to each well of the deepwell plates.

Plates were incubated at 30°C, 250 rpm overnight. When the OD600 nm reaches a value

comprised between 2-6, it means that the cells are in exponential phase. At this stage, the

medium was changed to promote induction. Cells were centrifuged at 4000 rpm for 10 min at

room temperature and the pellet was resuspended in 1 mL of BMMY medium supplemented

with PTM4 (1 µl for 1 mL). Deepwell plates were shaken at 200 rpm, 20°C during 3 days,

and 3% (v/v) of methanol was added every day. After 3 days of induction, cells were

centrifuged and the supernatant was transferred to a new deepwell and purified by the robot

using an in-house Tecan system as described in (Haon et al. 2015).

II.2.5. Recombinant production of recombinant enzymes

PaLPMO9H (protein ID CAP 61476) and variant without CBM were produced in Pichia

pastoris in flasks(Bennati-Granier et al. 2015). 5 mL of autoclaved YPD medium was added

in a tube and inoculated with a loop of Pichia cells from the Petri dish under agitation at 160

rpm at 30°C for four hours. The cultures for the production of recombinant proteins were

48

done at 30°C and 200 rpm. The medium used is BMGY medium (3.4 g L-1

Yeast Nitrogen

Base, without ammonium, 10 g L-1

of ammonium sulfate, 10 g L-1

glycerol, 10 g L-1

yeast

extract , 20 g L-1

peptone and 100 mL of phosphate buffer 1M pH 6). The time of incubation

was 16 hours until the OD600nm reached a value between 2 and 6. After centrifugation at

4000 rpm for 15 minutes at room temperature, the cell pellet was resuspended in 1/5 of

medium BMMY containing methanol (the same medium BMGY but without glycerol and

with 3% of pure methanol). The induction of the production of the enzyme was done at 30°C

during three days under constant agitation of 200 rpm and by the addition of 3% methanol

every day during three days.

II.2.6. Enzyme purification

After harvesting cells by centrifugation (2,700 g for 5 min, 4°C), the supernatant was adjusted

to pH 7.8 just before purification and filtrated on 0.22 µm membranes (Millipore, Molsheim,

France) The filtrate was loaded onto a 5 mL HiTrap HP column (GE Healthcare, Buc, France)

equilibrated with buffer A (Tris-HCl 50 mM pH 7.8, NaCl 150 mM, imidazole 10 mM) that

was connected to an Äkta purifier 100 (GE Healthcare). Each (His)6-tagged recombinant

enzyme was eluted with buffer B (Tris-HCl 50 mM pH 7.8, NaCl 150 mM, imidazole 500

mM). Fractions containing recombinant enzymes were pooled and concentrated with a 10-

kDa vivaspin concentrator (Sartorius, Palaiseau, France) and dialyzed against sodium acetate

buffer 50 mM, pH 5.2. The concentrated proteins were incubated with an equimolar

equivalent of CuSO4 overnight in cold room and buffer exchange in 50 mM sodium actetate

buffer pH 5.2 to remove the excess of CuSO4.

II.3. Protein analysis

II.3.1. Dosage of protein

The protein concentrations were determined by adsorption at 280 nm using a Nanodrop ND-

2000 spectrophotometer (Thermo Fisher Scientific) with theoretical molecular weights and

molar extinction coefficient derived from the sequences.

II.3.2. Protein electrophoresis

Proteins were loaded onto 10% Tris-glycine precast SDS-PAGE gels (BioRad, Marnes-la

Coquette, France) and stained with Coomassie Blue. The molecular mass under denaturing

conditions was determined with PageRuler Prestained Protein Ladder (Thermo Fisher

Scientific, IL, USA).

49

II.3.3. ICP-MS analysis

The ICP-MS analysis was performed as described in (Couturier et al. 2018) to quantify the

protein copper content. The samples were mineralized, then diluted in ultrapure water, and

analyzed by an ICAP Q apparatus (Thermo Electron, Les Ullis, France). The copper

concentration was determined using Plasmalab (Thermo Electron) software, at m/z=63.

II.4. Amplex red assay

A fluorimetric assay based on Amplex red and horseradish peroxidase was used as described

previously (Isaksen et al. 2014; Kittl et al. 2012). Measurements were carried out at 30°C in a

total volume of 100 µL using 50 mM phosphate buffer pH 6.0, 50 µM Amplex red (Sigma-

Aldrich, Saint-Quentin Fallavier, France), 7.1 U mL-1

horseradish peroxidase and 50 µM of

reductant (ascorbate, gallic acid, syringic acid, epigallocatechin, menodione, caffeic acid,

vanillic acid, p-coumaric acid, sinapic acid, 3-hydroxy anthranilic acid or L-cysteine). The

LPMO enzyme was added at a final concentration between 10 and 40 µM. Changes in

fluorescence were recorded with a Tecan Infinite M200 plate reader (Tecan, Männedorf,

Switzerland) over 24 min using excitation and emission wavelengths of 560 and 595 nm,

respectively. The specific activity was counted from H2O2 calibration curve, and the slope

(13,227 counts μmol−1

) was used to convert the fluorimeters’ readout (counts min−1

) into

enzyme activity.

II.5. Cellulose Binding Assays

The reaction mixtures were carried out at 0.3% (w/v) insoluble substrate loading (BMCC;

NFC; PASC) and 30 µg of proteins were added. The reactions were done in 50 mM sodium

acetate buffer pH 5.2 in a 200 µL final volume without any electron donor addition. The tubes

were incubated on ice for one hour with gentle mixing every ten minutes. After centrifugation

at 14,000 g for 10 min, the supernatant (containing the unbound proteins) was carefully

removed. Then, the polysaccharide pellets were washed twice (wash 1 and wash 2) by

resuspending in buffer and centrifuged at 14,000 g for 10 min. This step was repeated twice.

The remaining pellet was finally resuspended in SDS-loading buffer without dye (with a

volume equivalent to the unbound fraction removed) and boiled for 10 minutes to dissociate

any bound protein. Unbound, wash 2 and bound fractions (45 µL supplemented with 5 µL of

β-mercaptoethanol) were analyzed by SDS-PAGE to detect the presence or the absence of the

protein. The supernatant was recovered (supernatant 2: bound fraction). Forty five µL of

50

supernatant 1 (unbound fraction), wash 2 and supernatant 2 (bound fraction) were analyzed by

SDS-PAGE to detect the presence or the absence of the protein. A control sample without any

substrate was done in order to compare the results.

II.6. Enzyme assays

All the cleavage assays (300 μL final volume) contained 0.1 % (w/v) of substrate (PASC,

BMCC, NFC) 4.4 µM of PaLPMO9s, and 1 mM of L-cysteine, in 50 mM sodium acetate

buffer pH 5.2. The enzymatic reactions were incubated in a thermomixer (Eppendorf,

Montesson, France) at 50°C and 850 rpm for 16 hours. At the end of the reaction, the samples

were boiled at 100°C for 15 min and then centrifuged at 15,000 g for 10 min to separate the

soluble and insoluble fractions. Assays at 1% (w/v) PASC concentration were also done with

the previously mentioned conditions.

II.6.1. Combined assays

The LPMO enzymatic assays were carried out sequentially with a cellobiohydrolase from T.

reesei (CBH-I) as described in (Filiatrault-Chastel et al. 2019). Assays were performed in a

total volume of 800 µL containing 0.1% (v/w) cellulose in 50 mM pH 5.2 acetate buffer with

8 µg of LPMO enzyme and 1 mM L-cysteine. The samples were incubated in triplicate in a

thermomixer (Eppendorf) at 45°C and 850 rpm, for 24 h. The samples were then boiled for at

least 10 min and centrifuged at 15,000 g for 10 min. The supernatant was removed, and the

remaining insoluble fraction of the substrate was washed twice in buffer. Hydrolysis by CBH-

I (0.8 µg) was performed in 800 µL of 50 mM pH 5.2 acetate buffer for 2 h at 45°C and 850

rpm. The soluble fraction was analyzed as described below.

II.6.2. Analysis of oligosaccharides

Oxidized and non-oxidized cello-oligosaccharides generated after LPMO action were

analyzed by high-performance anion-exchange chromatography coupled with pulsed

amperometric detection (HPAEC-PAD) (ThermoFischer Scientific, USA) using a

CarboPacTM

PA1 column (2x250 mm) and CarboPacTM

PA1 guard column (2x50 mm) and a

flow rate of 0.25 mL min-1

as described by (Bennati-Granier et al. 2015) Non-oxidized

oligosaccharides were used as standards (Megazyme, Wicklow, Ireland).

51

II.7. Analysis of the insoluble fraction

The kraft fibers (100 mg) were adjusted to pH 5.2 with acetate buffer (50 mM) in a final

reaction volume of 20 mL. L-cysteine was present at a concentration of 1 mM of enzyme.

Purified LPMO enzyme was added to the substrate at a final concentration of 1.6 µM of kraft

fibers. Enzymatic incubation was performed at 50°C under mild agitation for 16 h. Samples

were then dispersed with a Polytron PT 2100 homogenizer (Kinematica AG, Germany) for

3 min and ultrasonicated with a QSonica Q700 sonicator (20 kHz, QSonica LLC., Newtown,

USA) at 350 W ultrasound power for 3 min. The reference sample was submitted to the same

treatment but it did not contain the LPMO enzyme.

II.7.1. Optical microscopy

Kraft fibers (reference and LPMO-treated) were deposited onto a glass slide and observed by

a BX51 polarizing microscope (Olympus France S.A.S.) with a 4X objective. Images

(N ≥ 20) were captured by a U-CMAD3 camera (Olympus, Japan). The concentration of the

fibers used was 2.5 g L-1

to visualize individual and separated fibers.

II.7.2. Atomic force microscopy (AFM)

Fiber dispersions were diluted at 0.1 g L−1

. Samples were dialyzed against ultrapure water

(spectral por-; molecular porous membrane tubing 12-14 kDa) for three days to remove

buffer, cysteine and released soluble sugars. They were later deposited onto mica substrates,

allowed to settle for 45 minutes and dried with Whatman filter paper. The final drying step

was done in the incubator at 40°C for ten minutes before passing them to AFM.

Topographical images on mica were registered by an Innova AFM (Bruker). The images were

collected in tapping mode under ambient air conditions (temperature and relative humidity)

using a monolithic silicon tip (FESPA-V2) with a spring constant of 2.8 N m−1

, and a nominal

frequency of 75 kHz. Image processing was performed with the WSxM 4.0 software.

A series of reference images (between 3 and 11) were recorded in order to ensure the

homogeneity of the sample.

II.7.3. Transmission electron microscopy (TEM).

Cellulose suspensions in water were deposited on freshly glow-discharged carbon-coated

electron microscope grids (200 mesh, Delta Microscopies, France) and the excess water was

removed by blotting. The sample was then immediately negatively stained with

52

phosphotungstic acid solution (1%, w/v pH 6) for 2 min and dried after blotting at room

temperature just before observation. The grids were observed with a Jeol JEM 1230 TEM at

80 kv. (Villares et al. 2018)

II.7.4. NMR analysis of the NFC samples: Sample preparation

All samples are centrifuged in order to concentrate cellulosic fibers: 4000 g for 15 min for

Samples and 4000g during 15 min followed by 8000 g during 30 min for NFC CTP sample.

Then cellulosic suspension are put into NMR 4mm rotor.

NMR conditions: Solid state 13

C CP/MAS NMR experiments were performed on an Avance

400 spectrometer operating at a 13

C frequency of 100.62 MHz. A double resonance H/X

CP/MAS 4 mm probe was used. The samples were spun at a rate of 9 kHz at room

temperature. The cross polarization pulse sequence parameters were: 3,5 μs proton 60° pulse,

3-5 ms contact time at 67.5 kHz, and 2 s recycle time. Typically, the accumulation of 30K to

100K (17h to 2,5 days) was used. Spectra were referenced using the carbonyl signal of

glycine at 176.03 ppm. All spectra obtained were processed and analysed using Bruker

Topspin version 3.2. Fitting of NMR spectra was performed using PeakFit software according

to the procedure described in (Villares et al. 2017).

II.7.5. Fourier transform infrared spectroscopy (FTIR)

Infrared spectra were obtained from KBr pellets containing freeze-dried cellulose samples

placed directly in a Nicolet iS50 FTIR spectrometer (Thermo Scientific) in absorbance mode.

All spectra were collected with a 4 cm-1

resolution after 200 continuous scans from 400 to

4000 cm-1

.

II.7.6. Neutral sugar composition

Identification and quantification of neutral sugar composition of cellulose samples were

performed by gas-liquid chromatography (GC) after sulfuric acid degradation (Hoebler et al.

1989). Five mg of dried cellulose were dispersed in 13 M sulfuric acid for 1h at 30 °C and

then hydrolyzed in 1 M sulfuric acid (2 h, 100 °C). Sugars were converted to alditol acetates

according to Blakeney et al. (Blakeney et al. 1983) and chromatographed on a TG-225 GC

column (30 × 0.32 mm ID) using a TRACE™ Ultra Gas Chromatograph (Thermo

Scientific™; temperature 205 °C, carrier gas H2). Standard sugars solution and inositol as

internal standard were used for calibration.

53

II.7.7. Colorimetric assay

The main objective of the assay developed by wang (Wang et al. 2018) is to quantify the

activity of C1-oxidizing LPMO on the insoluble fraction of the substrate. The main principle

of this method is the determination of the binding of cations (Ni2+

) to carboxyl groups formed

by the action of C1-oxidizing LPMOs on polysaccharides. The samples containing the

substrates were incubated with the enzymes. After incubation, the pellets were washed six

times with HEPES 10 mM buffer to remove all traces of enzyme, cysteine or charges that may

bias the results. The only detectable factor should be the oxidations created by the LPMO. 700

µL from each tube were removed from the supernatant and 540 µL of ethanol/HEPES 10 mM

were added. Samples were vigorously shaken and left undisturbed for 2 minutes. Then, 93 µL

of NiCl2 in ethanol/HEPES at a concentration of 2 mM were added and centrifuged after 2

min at 14000g for 5 minutes. 500 µL of the supernatant were mixed with pyrocatechol violet

(500 µl at 80 µM) and the absorbance was measured at a wavelength of 650 nm.

Control samples were prepared by adding 93 µL of NiCl2 (2mM) solution in ethanol/HEPES

and 840 µL of ethanol/HEPES 10 mM. After centrifugation (14000g for 5 min), 500 µL of

supernatant are added to 500 µL of pyrocatechol violet (80 µM). The Ni2+

ions present on the

fibers and bound to oxidations were detected by substracting total Ni2+

present in the control

containing no fibers from the Ni2+

found in the supernatant.

II.7.8. High Performance Size exclusion Chromatography coupled with Multi

Angle Light Scattering detector (HPSEC-MALLS)

II.7.8.1. Enzymatic treatment

The kraft fibers (100 mg) were adjusted to pH 5.2 with acetate buffer (50 mM) in a final

reaction volume of 50 mL. L-cysteine was present at a concentration of 1 mM. Purified

LPMO enzyme was added to the substrate at a final concentration of 10 mg per gram of kraft

fibers. Enzymatic incubation was performed at 50°C under mild agitation for 16 h. Later,

samples were boiled for 15 min. Half of the samples were then dispersed with a Polytron PT

2100 homogenizer (Kinematica AG, Germany) for 3 min and ultrasonicated with a QSonica

Q700 sonicator (20 kHz, QSonica LLC., Newtown, USA) at 350 W ultrasound power for 3

min. The rest of the samples were not subjected to mechanical treatment. The reference

sample was submitted to the same treatment but it did not contain the LPMO enzyme.

54

II.7.8.2. Fiber dissolution

Fiber dissolution was performed by solvent exchange by filtration. Fibers were firstly filtered

through 0.45 μm PTFE membranes to remove buffer and enzymes. The fiber cake was then

redispersed 3 times in anhydrous methanol (50 mL each time) followed by three additional

redispersions in anhydrous dimethlyacetamide (50 mL). Then the DMAc fibers swollen cake

was added to 5 or 10 mL of DMAc/LiCl (9% w/w) under mechanical stirring during 24 hours

before 10-fold dilution with anhydrous DMAc(Dupont and Mortha 2004).

II.7.8.3. Chromatographic analysis

Solutions were then filtered and injected on a size exclusion chromatography system

(OMNISEC Resolve, Malvern) with N,N-dimethylacetamide/lithium chloride (0.9 % w/v) as

the eluent. The SEC columns used were Viscoteck Tguard, LT4000L, LT5000L and

LT7000L. The system was equipped with a multi-angle laser light scattering Malvern SEC-

MALS 20 and OMNISEC Reveal devices (Malvern). Calculations were performed with a

dn/dc value of 0.136 mL g-1

and performed using OMNISEC software.

Polymethylmethacrylate PMMA60 was used as calibration standard and

Polymethylmethacrylate PMMA95 was used as verification standard.

II.7.9. Quartz crystal microbalance with dissipation monitoring (QCM-D)

The QCM-D measurements were performed with a Q-Sense E4 instrument (AB, Sweden)

using a piezoelectric AT-cut quartz crystal coated with gold electrodes on each side (QSX301,

Q-Sense). All measurements were carried out at 40 °C using the QCM flow cell modules.

Frequency (Δfn/n) and dissipation (ΔDn) changes were simultaneously registered at 5 MHz

fundamental resonance frequency and its several overtones as a function of time. Any

material adsorbed on the crystal surface induces a decrease of the resonance frequency (Δf). If

the adsorbed mass is evenly distributed, rigidly attached and small compared to the mass of

the crystal, f is directly proportional to the adsorbed mass per surface unit (Γ) using the

Sauerbrey’s equation (Sauerbrey 1959):

∆Γ = − C∆𝑓

𝑛 (1)

where C is the constant for the mass sensitivity of the quartz crystal (0.177 mg m-2

Hz-1

at f0 =

5 MHz) and n is the overtone number. The adsorption curves for multiple overtones were

55

used for estimating the adsorbed mass by the software QTools 3.1.25.604 (Q-Sense). Results

are expressed as the mean of at least 3 experiments.

Gold-coated quartz crystals were cleaned in piranha solution H2SO4/H2O2 (7:3, v/v), rinsed

exhaustively with Milli-Q water, and dried under a stream of nitrogen. Prior to use, QCM-D

quartz sensors were subjected to a plasma etching device (Harrick Plasma). Surfaces of

cellulose were prepared by the spin-coating method. Cellulose dispersions were dropped on

the gold electrodes and, after 5 min of adsorption, accelerated at 180 rpm s-1

to 3600 rpm for

60 s.

For the QCM-D experiments, a baseline was first established by continuously flowing acetate

buffer (50 mM) on the quartz crystal surface, then frequency and dissipation signals were off-

set to zero just before injection of LPMO in a continuous mode at a flow rate of 0.1 mL min-1

.

A rinsing step of the surface with acetate buffer (50 mM) was performed.

56

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B, Chaspoul F, Frandsen KE et al. . 2018. Lytic xylan oxidases from wood-decay

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Dupont AL, and Mortha G. 2004. Comparative evaluation of size-exclusion chromatography

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Hoebler C, Barry JL, David A, and Delortlaval J. 1989. Rapid acid hydrolysis of plant cell

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Isaksen T, Westereng B, Aachmann FL, Agger JW, Kracher D, Kittl R, Ludwig R, Haltrich

D, Eijsink VGH, and Horn SJ. 2014. A C4-oxidizing Lytic Polysaccharide

Monooxygenase Cleaving Both Cellulose and Cello-oligosaccharides. Journal of

Biological Chemistry 289(5):2632-2642.

Kittl R, Kracher D, Burgstaller D, Haltrich D, and Ludwig R. 2012. Production of four

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Villares A, Moreau C, Bennati-Granier C, Garajova S, Foucat L, Falourd X, Saake B, Berrin

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CELLULASE SUBSTRATES. Methods in Enzymology 160:19-25.

Chapter III

Preparation and characterization of the substrates

and the enzymes

59

III.1 Characterization of substrates used in this study:

In this thesis, we focused on four different cellulosic substrates (PASC), bacterial

microcrystalline cellulose (BMCC), nanofibrillated cellulose (NFC), and Kraft pulp fibers.

The substrates were chosen to dispose of the different structural patterns and hierarchical level

of organization of cellulose. Hence, PASC consists of amorphous cellulose with a DP max of

60 (Stålbrand et al. 1998). BMCC are rigid, rod-like cellulose nanocrystals that represent a

highly crystalline substrate. NFC contains both crystalline and amorphous regions at the

nanometric scale. Finally, Kraft pulp fibers represent the whole cellulose fiber containing both

crystalline and amorphous cellulose as well as a low percentage of hemicelluloses.

PASC was prepared from Avicel cellulose and it was not further characterized since it is a

common substrate widely described in literature (Bennati-Granier et al. 2015).(Zhang et al.

2006)

III.1.1. Bacterial Microcrystalline Cellulose (BMCC)

BMCC was obtained by acid hydrolysis of bacterial cellulose, which solubilizes the

amorphous regions of cellulose and allows the isolation of the rod-like cellulose nanocrystals.

Hydrolysis was performed with hydrochloric acid, resulting in neutral cellulose nanocrystals.

Differently from the hydrolysis with sulfuric acid, where nanocrystals have charged sulfate

groups on the surface that stabilizes the dispersion by electrostatic repulsions, BMCC were

rather neutral and nanocrystals tended to interact and aggregate. Therefore, the first

characterization was the determination of the Z-potential as a measure of the colloidal

stability of the BMCC dispersions. We obtained a Z-potential of -19.7 mV, which is

comparable to values reported in literature for cellulose nanocrystals prepared by acid

hydrolysis (Cherhal 2015). The average Z-size obtained by dynamic light scattering (DLS)

was 318.2 nm ± 76.9. Figure 19 shows the transmission electron microscopy (TEM) images of

bacterial cellulose nanocrystals. The images show the rod-like crystals of nanometric

dimensions. Nanocrystals are not highly aggregated, as the Z-potential values suggested. A

detailed analysis of the TEM images using Image J software revealed an average length of

855±215 nm and an average width of 17±5 nm (Figure 19) A thickness of 7 nm was

determined by atomic force microscopy (AFM), as previously observed for bacterial cellulose

nanocrystals (Kalashnikova et al. 2011) .(Kalashnikova et al. 2013)

60

Figure 19 TEM images of stained bacterial microcrystalline cellulose.

Figure 20 shows the Fourier-transform infrared (FTIR) spectrum of BMCC. The spectrum

reveals typical peaks corresponding to cellulose structural characteristics. (Li and Renneckar

2011; Lourdin et al. 2016; Oh et al. 2005).

Figure 20 Infrared spectrum of BMCC substrate

Figure 20 displays the FTIR spectra of the BMCC sample. The region from 3750 to 3000 cm-1

corresponds to stretching vibrations of the hydroxyl groups. The main peak at 3352 cm-1

is

attributed to intramolecular hydrogen bonds O(3)H…O(5) parallel to the β1-4 glycosidic

A B

D 25k15kCC D

61

bond. The region from 3000 to 2800 cm-1

is ascribed to the stretching vibrations of the C-H

bonds. The main band at 2902 cm-1

is assigned to the ring-CH of crystalline cellulose. Then,

the regions from 1500 to 1200 cm-1

and from 780 to 400 cm-1

correspond to bending

vibrations of the hydroxyl groups. The peaks at 1330 and 1363 cm-1

are respectively assigned

to –OH in plane bending and CH2 wagging motion (Carrillo et al. 2004); (Nelson and

O'Connor 1964);(Schwanninger et al. 2004). The peak at 1428 is attributed to the bending of

the O-H bond and the -CH2 symmetric bending or scissoring motion. Finally, the bands from

1100 to 780 cm-1

are assigned to the stretching vibrations of the C-O bond. The peak near

1110 cm-1

is attributed to the ring asymmetric stretching.

III.1.2.Nanofibrillated cellulose (NFC)

Nanofibrillated cellulose was provided by the Centre Technique du Papier (CTP, Grenoble,

France); therefore, several characterizations were performed including dry weight

measurement, optical microscopy, TEM, GPC, NMR and FTIR in order to choose one

substrate for the rest of the study.

III.1.2.1.Dry weight measurement:

The first characterization of the NFC samples was the determination of their concentrations.

For this purpose, we measured the dried mass of a known volume of solution. Table 2 reviews

the concentrations of the three samples studied.

Table 2: Average dry weight measurement of the three different types of NFC samples.

Sample Average of dry weight

(%) Standard deviation

Ex Piano 10% 10.77 0.10

Ex Piano 2% 2.00 0.08

NFC 2.5% 2.52 0.23

The average of dry weight of the three samples were different (Table 2). Ex piano 10% was

more viscous/concentrated than Ex piano (2%) and MFC (2.5%). Standard deviation values

are low and prove the accuracy of the average dry weight values.

62

III.1.2.2.Optical microscopy:

NFC samples were visualized by optical microscopy to obtain more information about their

aspect and morphology. Figures 21-23 review the optical microscopy images of the three

samples studied (Ex Piano 2 and 10% and NFC 2.5% CTP).

Figure 21 Optical microscopy image of Ex Piano (2%) at different scales.

In this Ex piano (2%) NFC sample big fibers (Figure 21 A and B) are approximately 200 µm

long and 40 µm width. It should be mentioned here that it is difficult to estimate the length

since the big fibers might not fit completely inside the microscopy and they were intermingled

with nanometric fibers (Figure 21 C and D).

63

Figure 22 Optical microscopy image of Ex Piano (10%) at different scales.

In the Ex piano (10%) NFC sample, big fibers (Figure 22 A and B) are approximately 400 µm

long and 75 µm width and were intermingled with nanometric fibers (Figure 22 C and D).

Similar observations were observed in the previous sample: Ex piano 2%.

A B

C D

64

Figure 23 Optical microscopy image of NFC (2.5%) CTP at different scales.

In the NFC 2.5% CTP sample, big fibers of approximately 250 µm to 750µm long and 25µm

width were found intermingled with nanometric fibers (Figure 23).

Both Ex Piano samples showed a higher content of intact cellulose fibers while the CTP

samples seemed more fibrillated, which meant that in the first and second samples, the

mechanical and the enzymatic treatments were not efficient enough to transform all the

cellulose fibers into micro and nanofibrillated cellulose. As for NFC CTP sample, fibers were

rarely found. This means that the third sample of MFC has been subjected to a more efficient

treatment yielding an aspect of highly entangled and mostly homogeneous microfibers of

cellulose.

III.1.2.3.Transmission electronic microscopy (TEM):

Optical microscopy images pointed at the presence of a nanometric fraction of cellulose not

detected by the microscope. Therefore, we visualized the samples by TEM in order to have a

view at the nanometric scale. Figures 24-26 show the TEM images of negatively stained

A B

C D

65

samples.

Figure 24 TEM microscopy of ex piano 2%.

TEM microscopy of ex piano 2% shows a web of short aggregated fibers (Figure 24 A, B and

D) and long intermingled fibers that are approximately 50 nm thick (Figure 24 C).

66

Figure 25 TEM microscopy of ex piano 10%

The fibers observed under the microscope looked drier than the previous samples since they

are more concentrated with less water content. The fibers are also more compact with smaller

fiber protruding all along the bigger fibers (Figure 25 A and Figure 25 B).

A B A B

D

12k 12k

C

TEM microscopy of ex piano 10% at a magnification of 50x

67

Figure 26 TEM microscopy of nanofibrillated cellulose 2% (NFC CTP)

TEM microscopy of nanofibrillated cellulose 2% shows a web of long intermingled fibers that

are approximately 50 nm thick (Figure 26 A to D).

In all cases, the TEM images showed fibers of nanometric dimensions, which confirmed the

fibrillation.

III.1.2.4. Carbohydrate composition:

The preparation of nanofibrillated cellulose involves the use of enzymes, for creating rupture

points in the fiber, and the mechanical delamination for separating the nanofibers. Generally,

other plant cell wall components, such as lignin, hemicelluloses or pectin are removed during

fibrillation; however, in most of the cases, a percentage of hemicelluloses closely bound to

cellulose remain. In order to determine this hemicellulose concentration in the samples, the

sugar content of the NFCs was determined by gas chromatography after acid hydrolysis

(Blakeney et al. 1983).

68

Table 3: weight % of sugars in the three samples of nanocelluloses.

Ex piano 2% Ex piano 10% NFC

Weight % Mean SD Mean SD Mean SD

Rha 0.3 0.1 0.4 0.1 0.4 0.0

Fuc 0.0 0.0 0.0 0.0 0.0 0.0

Ara 0.2 0.0 0.3 0.0 0.3 0.1

Xyl 1.7 0.1 1.7 0.1 1.7 0.1

Man 0.6 0.0 1.2 0.6 1.0 0.2

Gal 0.0 0.0 0.0 0.0 0.1 0.1

Glc 45.1 6.5 49.1 4.3 50.1 5.5

Total (cor.) 47.9 6.6 52.6 4.7 53.6 5.5

Sugar composition gave a yield of ≈50% for all samples, which indicated high water content.

The three samples Ex piano 2%, Ex piano 10% and NFC have glucose representing 40 to 50%

of their weight. However, they show trace amounts of xylose and mannose (Table 3), which

suggested the presence of xylan and mannan repectively. We can confirm then that almost

only cellulose and very few amount of hemicellulose constitute our samples. The remaining

50% of the samples are most likely due to incomplete hydrolysis.

69

III.1.2.5.Solid state NMR:

The NMR spectra of the three samples studied showed the characteristic peaks of cellulose:

C1 is at approximately 105 ppm, C4 has two peaks between 80 and 90 ppm, C6 is between 60

and 70 ppm, and the C2,C3, C5 region is between 70 and 80 ppm (Villares et al. 2017). -the

peaks corresponding to hemicelluloses appear around 102 ppm, next to the C1 peak, and

between 70 and 82 ppm, next to the C4 region.

Figure 27 NMR full spectra of the three cellulosic substrates.

NMR spectra are typical of Cellulose I in cellulosic fibers.. So the peak intensities have more

or less the same relative intensity across the three samples. Moreover, the background signal

(noise) is very high (Figure 27).

(ppm)

C1 region

C2, C3, C5 region

C6 region

C4 region

CP-MAS 13C NMR spectra

70

Figure 28 NMR C1 peak deconvolution of the three cellulosic substrates.

ZOOM – C1 region

(ppm)

Region of hemicellulose(if present)

3

71

Figure 29 NMR C4 peak deconvolution of the three cellulosic substrates.

The analysis of the C1 and C4 regions showed that there is no significant xylan in the three

samples as the hemicellulose % is < 5% (5% for CTP and 2-3% for Piano). The peaks in the

hemicellulose region correspond mainly to noise (Figure 29). This confirms the

polysaccharide analysis. Almost pure cellulose constitutes the samples.

III.1.2.6.Infrared analysis (FTIR):

FTIR analyses were performed to obtain more insight into the structural features of the three

NFC samples. The spectra did not reveal any difference between the three samples studied

(Figure 30).

Hemicelluloseregion

ZOOM - C4 region

(ppm)

Crystalline part

amorphous part

Example (for comparison) of the C4 regionfrom Birch cellulosic fibers

20% xylane

72

Figure 30 Infrared spectra of the four samples of nanocelluloses

The infrared spectra of the three samples of nanofibrillated cellulose (NFC CTP, Piano 2%,

Piano 10%) show similar profiles compared to the BMCC substrate previously discussed

(Figure 20). Indeed, there is no difference between the three NFC substrates (NFC CTP, Piano

2% and Piano 10%)

From the characterizations performed for the three NFC samples, we decided to work with the

NFC CTP sample because it showed a better degree of fibrillation, and the sugar composition

did not reveal significant differences between samples.

.

III-2 Recombinant production of the enzymes

III.2.1 Introduction:

III.2.1.1. Podospora anserina

Among the filamentous fungi, the ascomycete Podospora anserina has been studied for

decades for its impressive array of CAZymes involved in both cellulose and hemicelluloses

breakdown (Couturier et al. 2016). The sequencing of P. anserina genome was carried out in

2008 (Espagne et al., 2008). Annotation of genes encoding CAZymes revealed a large

diversity of plant cell wall-targeting activities and one of the highest numbers of CBMs

73

among fungal genomes available at the time. Comparison of closely-related fungi such as

Chaetomium globosum and Neurospora crassa and with the industrial workhorse

Trichoderma reesei confirmed that P. anserina encodes a complete machinery for plant cell

wall conversion including some auxiliary activity (AA) enzymes potentially targeting

recalcitrant cellulose and lignin. Enzymes targeting cellulose, i.e., cellobiohydrolases (GH6,

GH7) and endoglucanases (GH5, GH12, GH45) are abundant in P. anserina. Its genome

contains more than 100 CBMs, among which 28 CBM1, specific of cellulose recognition. Its

genome also encodes 33 AA9 LPMOs (PaLPMO9), eight of which contain a family 1

carbohydrate binding module (CBM1)-targeting cellulose. The expansion in genes encoding

AA9s has been observed in many fungal genomes. This gene multiplicity raises the question

of the functional relevance at the organism level, i.e., functional redundancy or functional

diversification and/or adaptations to substrate. In the secretomes of P. anserina after growth

on biomass, seven AA9 LPMOs were identified, five of which present a CBM1 (Poidevin et

al. 2014). More generally, modular AA9 LPMOs bearing a CBM1 at their C-terminus are

often predominantly secreted by filamentous fungi under lignocellulolytic conditions (Berrin

et al. 2017) but the role of these modules attached to LPMOs is not clearly established. In

another study, P. anserina cultured on soybean hulls produced a diverse set of CAZymes with

a total of 20 AA9 LPMOs identified using proteomics in the secretome (Mäkelä et al. 2017).

III.2.1.2. Podospora anserina LPMO enzymes

Out of the 33 AA9 LPMO enzymes found in the genome of P. anserina, 7 were

biochemically characterized (Bennati-Granier et al. 2015; Bey et al. 2013). All these AA9

LPMOs were active on cellulose with varying degrees of activity. Using high performance

anion exchange chromatography (HPAEC) a higher release of oxidized cello-oligosaccharides

from cellulose was observed for PaLPMO9A, PaLPMO9E and PaLPMO9H, all of which

harbor a CBM1 module (Bennati-Granier et al. 2015). Although it was a good indication that

the CBM1 module played an important role, the influence of the CBM1 module on enzyme

activity was not further investigated.

III.2.1.3.The PaLPMO9H enzyme

Out of the 7 AA9 from P. anserina that were biochemically characterized, PaLPMO9H was

the most deeply investigated because it displayed a peculiar broad substrate specificity. The

enzyme was shown to cleave mixed-linkage glucans, xyloglucan and glucomannan (Fanuel et

al. 2017) as well as cello-oligosaccharides (Bennati-Granier et al. 2015). Mass spectrometry

74

analysis of the released products revealed that PaLPMO9H catalyzes C4 oxidative cleavage

of mixed-linkage glucans and mixed C1/C4 oxidative cleavage of cellulose, glucomannan and

xyloglucan (Bennati-Granier et al. 2015; Fanuel et al. 2017), In addition, (Villares et al. 2017)

investigated the action of the enzyme LPMO9H on cellulosic fibers: after enzymatic treatment

and dispersion, LPMO-treated fibers showed dispersion. Based on NMR studies and

microscopy, it appears that LPMO creates nicking points that trigger the disintegration of the

fibrillar structure with rupture of the chains and release of elementary nanofibrils.

In this thesis, we decided to investigate the contribution of the CBM1 module to the catalytic

function of AA9. For this purpose, we produced LPMO9H enzyme with and without the

CBM1 (chapter III) then tested the enzymes on the different cellulosic substrates with

different aspects of crystallinity assessing the impact of the enzyme on the soluble (chapter

IV) and insoluble (chapter V) cellulosic fractions.

III.2.1.4.The roles of CBMs in cellulose-acting enzymes

The roles of CBMs in glycoside hydrolase function have been widely explored (see (Gilbert et

al. 2013) for a review). Indeed, many glycoside hydrolases that attack the plant cell wall

contain non-catalytic CBMs, which were first identified in cellulases (Van Tilbeurgh et al.

1986). CBMs are grouped into three types: type-A CBMs bind crystalline ligands while types

B and C bind internal or terminal regions of polysaccharides, respectively. CBM1 is a type-A

CBM, which binds crystalline substrates using a planar surface (Kraulis et al. 1989). CBMs

not only target the enzymes to their substrates to promote catalysis (Couturier et al. 2011)

(Hervé et al. 2010)], but sometimes they can also modulate enzyme specificity (Cuskin et al.

2012). CBMs are devoid of catalytic activity, but some studies suggest they play a role in the

amorphization of cellulose through non-hydrolytic disruption of the crystalline structure of

cellulose (Arantes and Saddler 2010); (Bernardes et al. 2019). CBM1 appended to AA9

LPMOs may influence substrate binding, enzyme activity and/or regioselectivity, but the data

are scarce and reported observations are contradictory. For instance, deletion of the CBM1 of

NcLPMO9C had no effect on the degradation of PASC (Borisova et al. 2015), whereas

removal of the natural CBM from cellulose-active bacterial LPMOs abolished their activity

(Crouch et al. 2016).

75

III.2.2. Results

III.2.2.1. Heterologous production of PaLPMO9H with and without CBM1

In filamentous fungi, a significant cohort of AA9 LPMOs is appended to CBM1. In order to

gain insight into the contribution of the CBM1 to the catalytic function of AA9 LPMOs, we

selected PaLPMO9H as a model enzyme based on previous biochemical analyses (Bennati-

Granier et al. 2015; Fanuel et al. 2017; Villares et al. 2017).

PaLPMO9H is a modular enzyme with two domains containing an N-terminal catalytic AA9

domain (16-243) and a C-terminal CBM1 domain (271-307) (Figure 31). These two domains

are connected through a serine/threonine/asparagine-rich linker comprising 27 amino acid

residues (Figure31). The gene was codon optimized for expression in Pichia pastoris

(Supplementary Figure S1) by insertion into the pPICZαA expression plasmid. A His tag was

added at the end of the protein in order to easily purify it.(Figure 31).

Figure 31 Schematic representation of the enzymes used in this study. LPMO-FL (full

length) and LPMO-CD cut at three different regions on the linker (catalytic domain)

with amino-acid numbering of the limits of each domain.

When the PaLPMO9H enzyme was truncated right after the catalytic module at position 244,

we were unable to successfully produce the corresponding recombinant protein in P. pastoris

(data not shown). Therefore, three other contructs with their CBM1 truncated by cutting at

three different regions on the linker (position 259; 267; 273) and thus differing in the length

of their linkers were also heterologously expressed by using synthetic genes (Figure 31)

76

(Supplementary Figure S1). Two types of Pichia competent cells were used (X33 and

Superman for less glycosylation) . Six transformants of each construct were produced and

purified by using the robot and run on a SDS-PAGE gel (Figure 32) to select the best

expressed protein with the highest intensity band. Following induction of selected P. pastoris

transformants, all the P. anserina AA9s (except IX5; IIIX1; IS3; IIIS2) were successfully

produced and purified to homogeneity (Figure 32).

Figure 32 Production of LPMO-CD in two different types of competent cells: X33(X)

and Superman(S). Construct I corresponds to the shortest construct cut at position 259

of the linker. Construct II corresponds to the intermediate construct cut at 267 of the

linker. Contruct III corresponds to the longest construct cut at 273 of the linker.

One out of the six transformants of each construct was chosen according to the intensity of the

protein band. The six chosen constructs were run on a gel (Figure 33).

77

Figure 33 Comparison of the sizes of the three different truncated LPMOs.

Electrophoresis analysis revealed that purified PaLPMO9s displayed apparent molecular

masses that were higher than the theoretical ones (Figure 33). This might be partly due to O-

and N-glycosylation that are predicted to be abundant in all of the PaLPMO9s, especially the

ones bearing serine/threonine-rich linker regions between the catalytic and the CBM1

modules that contain multiple glycosylation sites as already observed in other modular fungal

CAZymes. No clear differences were observed between recombinant LPMOs produced in

X33 and Superman.

Since the influence of the linker on the activity of LPMO was not the objective of the study

and since the shortest construct was successfully expressed, the LPMO with the shortest

linker was chosen (Construct I that was expressed in Superman competent cell: IS4) to be the

construct of LPMO-CD to be further studied throughout the thesis (Figure 33).

Therefore, we decided to leave 16 amino-acid residues of the linker to promote production of

the recombinant enzyme. Using this strategy, we successfully produced the CBM1-free

PaLPMO9H enzyme truncated at position 259. In the rest of the study, the PaLPMO9H with

the CBM1 is named LPMO-FL (full-length), and the PaLPMO9H without the CBM1 is

named LPMO-CD (catalytic domain). As expected, deletion of the CBM1 decreased the

enzyme’s molecular mass from ~38 kDa (LPMO-FL) to ~33 kDa (LPMO-CD). Apparent

15

25

35

40

5570100130180

78

molecular mass was still slightly higher than theoretical molecular mass (25.7 kDa) due to

predicted O- and N-glycosylations (Figure 33).

LPMOs are copper-dependent enzymes, which makes it crucial to check the correct copper

protein loading. The amount of copper in each enzyme was quantified using inductively-

coupled plasma mass spectrometry (ICP-MS). Both enzymes were equally loaded with ~one

copper atom per molecule (i.e. 10.3 and 10.8 µM of Cu2+

for 10 µM of LPMO-FL and

LPMO-CD, respectively).

In order to compare the activity of the active site of both enzymes in the absence of substrate,

a fluorimetric assay was performed to both enzymes. (Figure 34) The basic principle of this

test is that the hydrogen peroxide is produced by the active site of the enzyme and is induced

by an electron donor. The H2O2 activates horseradish peroxidase. The enzyme transforms

Amplex red to resorufin which can be detected by the spectrophotometer.

a

79

Figure 34 Fluorimetric assay for the generation of hydrogen peroxide by LPMO-FL (a)

and LPMO-CD (b). Reactions were run in triplicates;

Conclusion:

We managed to produce the LPMO-CD without its CBM module in Pichia pastoris.

Biochemically, the characteristics of both enzymes (LPMO-FL and LPMO-CD) are

comparable in terms of Copper content and Hydrogen peroxide production (Figure 34).

Therefore these enzymes will be suitable for further comparative enzymatic characterization

on different types of cellulosic substrates.

b

80

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Polysaccharide Monooxygenases (Family GH61) from Podospora anserina. Applied

and Environmental Microbiology 79(2):488-496.

Blakeney AB, Harris PJ, Henry RJ, and Stone BA. 1983. A simple and rapid preparation of

alditol acetates for monosaccharide analysis. Carbohydrate Research 113(2):291-299.

Borisova AS, Isaksen T, Dimarogona M, Kognole AA, Mathiesen G, Varnai A, Rohr AK,

Payne CM, Sorlie M, Sandgren M et al. . 2015. Structural and Functional

Characterization of a Lytic Polysaccharide Monooxygenase with Broad Substrate

Specificity. Journal of Biological Chemistry 290(38):22955-22969.

Carrillo F, Colom X, Sunol J, and Saurina J. 2004. Structural FTIR analysis and thermal

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Couturier M, Feliu J, Haon M, Navarro D, Lesage-Meessen L, Coutinho PM, and Berrin J-G.

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Couturier M, Tangthirasunun N, Ning X, Brun S, Gautier V, Bennati-Granier C, Silar P, and

Berrin J-G. 2016. Plant biomass degrading ability of the coprophilic ascomycete

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Crouch LI, Labourel A, Walton PH, Davies GJ, and Gilbert HJ. 2016. The Contribution of

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Cuskin F, Flint JE, Gloster TM, Morland C, Baslé A, Henrissat B, Coutinho PM, Strazzulli A,

Solovyova AS, and Davies GJ. 2012. How nature can exploit nonspecific catalytic and

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Fanuel M, Garajova S, Ropartz D, McGregor N, Brumer H, Rogniaux H, and Berrin J-G.

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Chapter IV

Depicting the activity of the LPMO enzyme by

analyzing the soluble fraction

85

IV.1 Absence of CBM1 alters LPMO cellulolytic activity at low

substrate concentration

The action of LPMO-FL was first evaluated on three different types of cellulose, i.e.

phosphoric acid-swollen cellulose (PASC), nanofibrillated cellulose (NFC), and bacterial

microcrystalline cellulose (BMCC) by the determination of soluble sugars released upon the

enzymatic action (Figure 35a). For the oxidized sugars, there were not standards available,

therefore, the peaks were assigned by comparison with literature (Bennati-Granier et al.

2015). As previously shown (Bennati-Granier et al. 2015), LPMO-FL released C4-oxidized

(C4ox), C1-C4-oxidized and non-oxidized oligosaccharides including cellobiose (Glc2),

cellotriose (Glc3), cellotetraose (Glc4), cellopentaose (Glc5) and cellohexaose (Glc6) from

PASC . However, using NFC as substrate led to less products released including Glc2, Glc3,

Glc4, some C4ox products and some slight peaks of C1-C4ox products. However, using a

more recalcitrant crystalline substrate (BMCC) led to barely detectable products including

Glc3 and Glc4 as the non-oxidized products and products oxidized at the C4 position (C4ox)

as the oxidized products (Figure 35a). BMCC is the most recalcitrant substrate having more

crystalline regions than the other substrates PASC and NFC. We then compared the action of

both LPMO-FL and LPMO-CD by measuring the release of sugars from PASC (Figure 35b).

LPMO-FL released higher amounts of soluble sugars (both oxidized and non-oxidized

oligosaccharides) compared to LPMO-CD where soluble sugars were barely detectable

(Figure 35b).

86

Figure 35 Analysis of soluble degradation products. a Products generated by LPMO-FL

upon degradation of 0.1% PASC, NFC or BMCC with 4.4 μM of LPMO in the presence

of 1 mM of L-cysteine, at 50 °C for 16 h. b Analysis of soluble degradation products

generated by LPMO-FL and LPMO-CD upon degradation of 0.1% PASC with 4.4 μM

of LPMO in the presence of 1 mM of L-cysteine, at 50 °C for 4 h.

10 15 20 25 30 35 40 45

0

200

400

600

800C

harg

e [

nC

]

Retention Time [min]

PASC

NFC

BMCC

PASC (no enzyme)

C4oxC4ox C1-C4ox

Glc

3

Glc

6

Glc

4

Glc

5

Glc

2

Glc2-Glc6

10 15 20 25 30 35 40 45

0

800

Charg

e [

nC

]

Retention Time [min]

LPMO-FL

LPMO-CD

No enzyme

200

400

600

Glc

2

Glc

3

Glc

6

Glc

4

Glc

5

C4ox C4ox C1-C4ox

Glc2-Glc6

a

b

87

IV.2 LPMO-FL and LPMO-CD are both able to cleave cellohexaose

Since LPMO-FL is active on Glc4, Glc5 and Glc6 soluble oligosaccharides (Bennati-Granier

et al. 2015), we investigated the activity of both LPMO-FL and LPMO-CD on cellohexaose

as substrate (Figure 36). A time-course analysis revealed that both enzymes were able to

cleave Glc6 (1 mM), leading mainly to Glc3 and Glc4 non-oxidized products with

respectively 190 µM and 140 µM of Glc4 for LPMO-FL and LPMO-CD after 24 hours of

incubation and approximately 25 µM of Glc3 for both enzymes after 24 hours of incubation.

Although LPMO-FL showed slightly better activity than LPMO-CD over the 24-h time-

course, the observed cleavage of Glc6 by LPMO-CD confirms that the enzyme lacking the

CBM1 module is still functional.

Figure 36 Time -course analysis and quantification of the Glc4 (plain lines) and Glc3

(dotted lines) released by LPMO-FL (triangles) and LPMO-CD (circles) acting on Glc6

(1mM) over a total period of 24 hours. Enzyme concentration was 1 µM.

0

50

100

150

200

250

0 5 10 15 20 25

pro

du

ct f

orm

atio

n (

µM

)

Time (hours)

88

IV.3 The CBM1 favors the binding of LPMO to cellulosic substrates

Since LPMO-CD showed some activity on the soluble sugar Glc6, binding to cellulose was

evaluated for both enzymes using a pull down assay. The aim of this experiment was to reveal

whether or not the cellulolytic activity of the LPMO enzyme is influenced by its binding

capacity with CBM1.

LPMO-FL and LPMO-CD binding to PASC, BMCC and NFC was assessed in the absence of

reductant using pull-down assays to evaluate the impact of the CBM1 (Figure 37). LPMO-FL

was observed in the bound fraction of all three cellulosic substrates tested (lane 4). This

observation means the LPMO-FL is able to bind to the three cellulosic substrates that differ in

terms of structure (crystallinity or amorphous). However, in the absence of CBM1, there were

no bands corresponding to LPMO-CD in the bound fraction. Although, pull down assays are

qualitative experiments, our results suggest that LPMO-FL bind more tightly to the cellulosic

substrates compared to LPMO-CD. In other words, these results suggest that CBM1 promotes

LPMO binding to the cellulosic substrates.

89

Figure 37 Qualitative cellulose binding assays. Binding of LPMO-FL and LPMO-CD to

(a) PASC (0.3% (w/v)), (b) NFC (0.3% (w/v)) and (c) BMCC (0.3% (w/v)). Lane 1,

control (no substrate); lane 2, unbound material; lane 3, wash and lane 4, bound

fraction. Experiments were carried out on ice using 30 µg of proteins and 50 mM sodium

acetate buffer pH 5.2 in 200 µl final volume without addition of L-cysteine. The ladder

size is indicated in kDa.

IV.4 Combined action of LPMO-FL and LPMO-CD with a

cellobiohydrolase

In order to assess the impact of LPMO-CD on cellulosic substrates, we assayed LPMO-FL

and LPMO-CD enzymes in combination with the reducing end-acting cellobiohydrolase

(family GH7 CBH-I) from T. reesei. Cellulosic substrates were sequentially pretreated with

either LPMO-FL or LPMO-CD before addition of the CBH-I enzyme. As both LPMOs and

CBH-I act on soluble substrates, we implemented an LPMO post-treatment washing step to

assess the impact of LPMO treatment only on the insoluble fibers. LPMO pretreatment was

beneficial on PASC and NFC but had no visible effect on the crystalline substrate BMCC

(Figure 38). Pretreatment with either LPMO-CD or LPMO-FL increased cellobiose release

from NFC substrate by approximately 30%. However, LPMO-FL pretreatment was more

efficient on PASC substrate (60% increase in cellobiose production) compared to the LPMO-

CD. Taken together, these results show that neither of the two LPMOs targets the crystalline

fraction of cellulose. We believe that both LPMOs target amorphous regions thus facilitating

1 2 3 4 1 2 3 4

c

LPMO-FL LPMO-CD

90

CBH-I activity on crystalline cellulose. Moreover, under these experimental conditions the

presence of the CBM1 module is not strictly required for LPMO action.

Figure 38 Combined action of LPMO-FL and LPMO-CD with a cellobiohydrolase

(CBH). The cellobiose released (in μM) from the three cellulosic substrates NFC, PASC

and BMCC was quantified using ion chromatography.

IV.5 Increasing substrate concentration reduces the need for the

CBM1

The next step was to assess whether substrate concentration has an influence on the activity of

the enzymes. We increased the substrate concentration to 1% (w/v) to foster the probability of

enzyme–substrate interactions in a CBM-free context. At high substrate concentration, soluble

sugars released by LPMO-CD became detectable and were in the same range as soluble

sugars released by LPMO-FL from PASC (Figure 39). Interestingly, C1-oxidized (C1ox)

products (Glc2ox-Glc4ox), which were barely detectable using LPMO-FL, were abundantly

released by LPMO-CD (Figure 39). The C4-oxidized products eluting at around 30 minutes

were less abundant whereas the C1/C4-oxidized products eluting between 41 and 42 minutes

were slightly increased. The absence of the CBM induced a modification of the

regioselectivity pattern of the enzyme (Figure 39).

91

Figure 39 Analysis of degradation products generated by LPMO-FL and LPMO-CD.

HPAEC chromatograms of the oligosaccharides released upon degradation of PASC

[1% (w/v)] with 4.4 μM of LPMO in the presence of 1 mM of l-cysteine, at 50 °C for 16

h. The sum of C1-oxidized (C1ox) and C4-oxidized (C4ox) oligosaccharides is indicated

in the inset. *Glc2ox co-eluted with Glc6

The functional relevance of CBMs and their contribution to the activity of the LPMO

enzymes have already been investigated (Crouch et al. 2016);(Danneels et al. 2018), but in

several cases, analysis surprisingly found modest and contradictory effects on enzyme

activity. The role of CBMs appended to glycoside hydrolases has been explored in depth

(Gilbert et al. 2013), and it is generally acknowledged that the presence of CBMs increases

the concentration of proteins at the surface of the substrate, thus increasing the activity of the

enzyme (Guillen et al. 2010). Removal of the CBM attached to cellulases dramatically

decreases the activity on insoluble substrates but not on soluble substrates (Kotiranta et al.

1999; Paakko et al. 2007; Stahlberg et al. 1991). A similar pattern was observed here with

92

PaLPMO9H, as loss of the CBM dramatically affected the release of soluble sugars from

cellulose whereas activity was retained on soluble cellooligosaccharides.

When the substrate concentration of cellulose was increased, the lack of CBM did not seem to

impede the action of PaLPMO9H (LPMO-CD), and soluble products were detected in the

same range as the full-length enzyme. A similar pattern of action was observed with cellulases

where a reduced amount of water counterbalanced the need for CBMs (Varnai et al. 2013).

Our results are in line with the hypotheses drawn by Courtade et al. (Courtade et al. 2018)on a

cellulose-active AA10 LPMO. Indeed, multiple cleavages are needed at the cellulose surface

to release enough soluble products that can then be detected by ion chromatography. Here, we

observed that the CBM1 appended to an AA9 LPMO promotes binding to cellulose and

anchors the enzyme to the substrate, facilitating multiple localized cleavages.

93

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Crouch LI, Labourel A, Walton PH, Davies GJ, and Gilbert HJ. 2016. The Contribution of

Non-catalytic Carbohydrate Binding Modules to the Activity of Lytic Polysaccharide

Monooxygenases. Journal of Biological Chemistry 291(14):7439-+.

Danneels B, Tanghe M, and Desmet T. 2018. Structural Features on the Substrate-Binding

Surface of Fungal Lytic Polysaccharide Monooxygenases Determine Their Oxidative

Regioselectivity. Biotechnology journal.

Gilbert HJ, Knox JP, and Boraston AB. 2013. Advances in understanding the molecular basis

of plant cell wall polysaccharide recognition by carbohydrate-binding modules.

Current Opinion in Structural Biology 23(5):669-677.

Guillen D, Sanchez S, and Rodriguez-Sanoja R. 2010. Carbohydrate-binding domains:

multiplicity of biological roles. Applied Microbiology and Biotechnology 85(5):1241-

1249.

Kotiranta P, Karlsson J, Siika-Aho M, Medve J, Viikari L, Tjerneld F, and Tenkanen M.

1999. Adsorption and activity of Trichoderma reesei cellobiohydrolase I,

endoglucanase II, and the corresponding core proteins on steam pretreated willow.

Applied Biochemistry and Biotechnology 81(2):81-90.

Paakko M, Ankerfors M, Kosonen H, Nykanen A, Ahola S, Osterberg M, Ruokolainen J,

Laine J, Larsson PT, Ikkala O et al. . 2007. Enzymatic hydrolysis combined with

mechanical shearing and high-pressure homogenization for nanoscale cellulose fibrils

and strong gels. Biomacromolecules 8(6):1934-1941.

Stahlberg J, Johansson G, and Pettersson G. 1991. A NEW MODEL FOR ENZYMATIC-

HYDROLYSIS OF CELLULOSE BASED ON THE 2-DOMAIN STRUCTURE OF

CELLOBIOHYDROLASE-I. Bio-Technology 9(3):286-290.

Varnai A, Siika-aho M, and Viikari L. 2013. Carbohydrate-binding modules (CBMs)

revisited: reduced amount of water counterbalances the need for CBMs.

Biotechnology for Biofuels 6.

Chapter V

Depicting the activity of the LPMO enzyme by

analyzing the insoluble fraction

97

To detect the activity of LPMO on the insoluble fraction, we firstly focused on the changes in

morphology of the fibers after the LPMO action, visualized by microscopic techniques, and

the variations in the cellulose chain length, evaluated by high performance size exclusion

chromatography (HPSEC). The mass release upon enzymatic action, in synergy with

cellobiohydrolase-I, was monitored in real time by quartz crystal microbalance with

dissipation. Finally, we also tried to quantify the oxidized groups formed after the action of

LPMO by a colorimetric assay.

V.1 Direct visualization of fiber morphology by microscopy

In an effort to gain more insights into the action of LPMOs and the role of the CBM, we

evaluated the changes in morphology of kraft fibers in response to incubation with LPMO

using optical microscopy and atomic force microscopy (AFM) techniques.

First, we investigated the fiber structure using optical microscopy. Original kraft fibers are

tens of micrometers in diameter and around 1 mm long (Figure 40a). After LPMO treatment,

there were no visible changes in physical appearance of the fibers, i.e. fibrous morphology or

dimensions, in LPMO-FL (Figure 40b) and LPMO-CD-treated samples (Figure 40c). As

previously described (Villares et al. 2017b), the action of LPMOs alone does not produce a

noticeable disintegration of kraft fibers, similarly to the action of cellulases (Henriksson et al.

2007; Nechyporchuk et al. 2015; Paakko et al. 2007). Therefore, after LPMO treatment, fibers

were mechanically dispersed and then subjected to a short ultrasound treatment. Dispersion

revealed the effect of LPMO on kraft fibers. Control samples showed some slight fibrillation

whereas both LPMO-treated samples showed clear cell wall delamination (Figure 40d-f).

Both LPMO-FL and LPMO-CD seemed to create weak points within the fiber that facilitated

the mechanical disintegration.

To get a better picture of the effect of the LPMOs, we used AFM to analyse samples (Figure

40g-i). A zooming on the fibers was done to obtain dimensions of 50 µm of the topographical

aspects of the fibers. Topography images showed the presence of large fibers in control

samples and a clear dissociation in LPMO-treated samples. LPMO-FL produced fibrillation of

kraft fibers, forming an entangled network of ~5 nm-diameter nanofibrils. LPMO-CD also

produced a network of disintegrated fibers, but with thicker fiber bundles. Comparing the

appearance of the fibers treated with LPMO-FL or LPMO-CD against controls, it is evident

that both enzymes influence the cohesion and architecture of the fibers, making them more

prone to the mechanical forces caused by the dispersion. Based on AFM images, both LPMOs

98

reduced fiber cohesion, but the presence of CBM appeared to enable LPMO-FL to defibrillate

cellulose.

3D images of the fibers without mechanical treatment were done (Figure 40 j-l). More AFM

3D images are provided in the supplementary data (Supplementary figure S2). The aim was to

visualize the direct effect of the enzymes LPMO-FL and LPMO-CD without any intervention

of the mechanical treatment. This can help in the detection of the factors in the topographical

aspect of the fibers which is causing the defibrillation after the mechanical treatment. The

control fibers showed a groove that was originally present (see Figure 40 j and supplementary

figure S2 (a-b)). The original kraft fiber was compact and had a cross section showing a slight

depression between two thicker shapes on the edges and a thickness of 3.8µm. The 3D image

of LPMO-FL treated fiber was hollow and filled with tearings. Only the thicker edges of the

fiber resisted the enzymatic action and remained after LPMO-FL treatment. It was of a

maximum thickness of 3.7 µm (Figure 40k and Supplementary figure S2(c-d)). LPMO-CD-

treated fiber had a groove in the middle but with less tearings than the LPMO-FL-treated

fiber. The surface in the middle remained smooth (Supplementary figure S2 (e-f)) and lower

than the edges but with less or no tearings. Additionally, one relevant observation is that the

probe of the AFM could easily access the LPMO-treated fibers but it is on its limit in the

control sample. This means that the control fibers are more compact than the enzymatically-

treated samples. More 3D images need to be done in order to have a statistically relevant

approximation of the topographical aspects of the fibers both enzymatically and non-

enzymatically-treated.

99

Figure 40 Morphology of LPMO-treated kraft fibers. Optical microscopy images of

kraft fibers before (a–c) and after (d–f) mechanical dispersion for control samples (a, d),

LPMO-FL-treated fibers (b, e) and LPMO-CD-treated fibers (c, f). AFM topographical

images after LPMO treatment and dispersion for control kraft fibers (g), LPMO-FL-

treated fibers (h) and LPMO-CD-treated fibers (i). AFM topographical 3D images with

LPMO treatment without mechanical dispersion: control fiber (j), LPMO-FL-treated

fiber (k), and LPMO-CD-treated fiber (l).

j k l

100

V.2.Evaluation of chain length by high performance size exclusion

chromatography (HPSEC):

V.2.1.The weight average molecular weight of the non-mechanically-

treated samples

The impact of the LPMO action on the cellulose chain length was evaluated by high

performance size exclusion chromatography (HPSEC) coupled to multi-angle laser light

scattering detection (MALLS). Since cellulose is insoluble, the first step of this determination

was the dissolution of cellulose fibers into dimethylacetamide (DMAc) mixed with LiCl.

(Dupont and Mortha 2004). For this purpose, a solvent exchange from water to ethanol and to

DMAc was performed. First the samples were filtered to remove the buffer and the enzyme.

Then, the fibers were activated into water under magnetic stirring for one hour. Water was

exchanged to methanol by filtration, and then exchanged to anhydrous DMAc. Samples were

agitated overnight then the fibers were filtrated and anhydrous DMAc was added again and

agitated for one hour. The fibers were then filtered and dispersed in DMAc/LiCl 8%.

For the non-mechanically-treated samples, two sets of samples were injected on the HPSEC

and each set was reinjected to verify the accuracy of the measurement. An average of both

sets and their reinjections is shown in Figure 41. The controls (with or without Cysteine)

showed a weight average molecular weight of approximately 900,000±70,000 g/mol. When

cellulose fibers were treated with both LPMOs a decrease of the molecular weight was

observed, and the effect was slightly more pronounced in the case of LPMO-CD

(590,000±225,000 g/mol) compared to LPMO-FL (674,000±514,000 g/mol). The decrease of

the molecular weight demonstrated the action of both LPMOs on the cellulose fibers.

Nevertheless, the standard deviation became higher in the LPMO-FL-treated sample, and it

was lower in the LPMO-CD-treated sample. This data clearly showed that the non-

mechanically-treated samples were not reproducible in their weight average molecular weight

(Mw). Thus, these data cannot be exploited and no conclusion can be drawn from them. One

probable reason for this non-reproducibility might be the non-efficiency of the dissolution of

the cellulosic kraft fibers. The fibers were probably not individualized enough for the

penetration of the DMAc/LiCl complex especially those that were not mechanically-treated.

Another reason might be the presence of water in the samples that might hinder the formation

of the DMAc/LiCl complex which dissolves the cellulosic fibers since both of them are

hygroscopic.

101

Figure 41 Mw average of the non-mechanically-treated samples

V.2.2.The weight average molecular weight of the mechanically-treated

samples

In the mechanically-treated samples (Figure 42), the average of two sets of samples was done.

The weight average molecular weight was in the range of 800,000 g/mol in the control

without Cysteine and 900,000 g/mol in the control with Cysteine. The LPMO-FL-treated

sample showed a decrease in weight average Mw as expected (to approximately 700,000

g/mol). LPMO-CD also shows a small decrease with respect to the control Cysteine (about

780,000 g/mol). The mechanically-treated samples showed higher reproducibility than the

non-mechanically-treated samples. Results pointed to similar decreases in molecular weight

for mechanically and non-mechanically treated samples. This assumption would indicate that

the decrease in chain length is produced by the action of LPMOs, and not by the mechanical

treatment.

102

Figure 42 Mw average of the mechanically-treated samples

The decrease of the molar mass upon the action of both LPMOs clearly demonstrated the

effect of these enzymes on the chain length. However, the decrease in the degree of

polymerization, or chain length, is not very pronounced, which is desired for maintaining the

mechanical properties of cellulose fibers.

V.2.3.The number average molecular weight and the polydispersity of

the non-mechanically-treated samples

The number-average molecular weight (Figure 43a) is highly dispersed and not reproducible

among the non-mechanically treated samples, especially in the LPMO-treated samples. No

conclusion can be drawn from the two sets #1 and #2. The polydispersity index (Mw/Mn)

increased upon the action of LPMOs (Figure 43b), which confirmed the release of lower

molecular weight chains. Nevertheless, the polydispersity index is not reproducible among

both sets of samples (Figure 43b). There-are four reasons for the irreproducibility of the

results: (i) variation of the recovery rate since the samples are not solubilized to the same

extend (ii) The loss of low masses during the solvent exchange process (iii) Dynamic light

scattering slightly sensitive to the small masses (iv) Difficulty on the integration since Mn

values are sensitive to low molar mass.

103

Figure 43 Number-average molecular weight and polydispersity index of set #1 and

set#2 of the non-mechanically-treated samples.

V.2.4.The number average molecular weight and the polydispersity of the

mechanically-treated samples

Similar conclusions can be deduced from the mechanically-treated samples (Figure 44). Both,

the number average molecular weight and the polydispersity index have high standard

deviation and no conclusions can be drawn. Normally, the number average molecular weight

and the weight average molecular weight should be extracted from the same data.

The probable reasons why HPSEC experiments did not give exploitable data are: (i) water has

entered the solutions. (ii) The solutions should be passed directly to analysis and not left to be

0

100000

200000

300000

400000

500000

600000

Control Control Cysteine LPMO-FL LPMO-CD

Nu

mb

er

ave

rage

mo

lecu

lar

we

igh

t (g

/mo

l)

Mn average of set#1 and set#2

0

5

10

15

20

25

30

35

40

Control Control Cysteine LPMO-FL LPMO-CD

Po

lyd

isp

ers

ity

ind

ex

Mw/Mn average of set#1 and set#2 b

a

104

settled down. Time can change the molecular weight distributions of the solutions. A solution

would be to repeat the same samples several times to have statistically relevant results.

Figure 44 Number-average molecular weight and polydispersity index of set #1 and

set#2 of the mechanically-treated samples.

V.3. Real-time monitoring of action of LPMO on regenerated cellulose

by quartz crystal microbalance with dissipation monitoring (QCM-D)

Sauerbrey (Sauerbrey 1959a) was the first to recognize the potential usefulness of the Quartz

Crystal Microbalance (QCM-D) technology and demonstrate the extremely sensitive nature of

these piezoelectric devices towards mass changes at the surface of QCM-D electrodes.

105

V.3.1.The QCM-D technique

The QCM-D technique is based on the properties of the quartz piezoelectric detectors

subjected to an electric field that induces a mechanical shear wave of the crystal. The

adsorption of any material on the surface of the quartz induces a shift of the resonance

frequency (∆fn) which permits to follow in real-time the adsorption of the polymers on the

quartz surface. Due to its high sensitivity, this technique has been used successfully to

monitor a wide range of adsorption processes. For instance, QCM-D has already been used to

study the adsorption of hemicelluloses and its derivatives (Kohnke et al. 2011; Villares et al.

2017a) (Villares et al. 2015) as well as other biopolymers (Villares et al. 2013).

If the adsorbed polymers are uniformely distributed and rigidly fixed, the Sauerbrey

equation can be used to quantify the mass adsorbed by the resonance frequency shift

(Sauerbrey 1959b): ∆Γ = − C∆𝑓

𝑛

Where C is the Sauerbrey constant: constant of the mass sensitivity of the quartz crystal

(0,177 mg.m-2

.Hz-1

) and n is the number of harmonics. 𝑛 = 1, 3, 5, 7, 9, 11 and 13.

The third harmonic (5 MHz) was used for the evaluation of the QCM-D data in this work.

The second piece of information that can be obtained from the QCM-D analysis is the

dissipation factor ΔD (Rodahl and Kasemo 1996). This value gives an indication of the

mechanical behavior (viscoelastic) of the adsorbed layer on quartz since it reflects how the

layer dissipates the energy of the oscillation waves of quartz. Indeed, the resonance frequency

of the quartz crystal depends on the total oscillating weight, including the coupled water. The

Figure 45 gives a schematic representation of the measurements of QCM-D with two types of

deposited layers, rigid and flexible. If the adsorbed layer is not fully elastic, friction losses

occur, leading to damping of the oscillation with greater amplitude. The change of the

dissipation factor ∆D is introduced. The dissipation factor (D) is defined as the ratio between

the energy dissipated during an oscillation period and the energy stored in a period of the

oscillating system.

𝐷 =𝐸𝐷𝑖𝑠𝑠𝑖𝑝𝑎𝑡𝑒𝑑

2𝜋𝐸𝑠𝑡𝑜𝑟𝑒𝑑

106

Where EDissipated and Estored are the total energy dissipated and stored during an oscillation

respectively. The energy dissipation factor therefore provides a measure of the stiffness or

viscoelasticity of the adsorbed layers (Rodahl and Kasemo 1996).

Figure 45 Schematic representation of the decreasing of the signal for the QCM-D

measurements on a rigid layer adsorbed on the surface with low dissipation, and a

flexible and viscoelastic layer with high dissipation.

V.3.2.Preparation of the regenerated amorphous cellulose layer

After the gold-coated quartz crystals have been cleaned and have been subjected to plasma

cleaner, surfaces of cellulose were prepared by spin coating: cellulose dispersions were

dropped on the gold electrodes, and, after 5 min of adsorption, accelerated at 180 rpms-1

to

3600 rpm for 60s.

V.3.3.Control experiment: Injecting cysteine or ascorbate and CBHI to the

regenerated amorphous cellulose surface

The experiments were carried on at a temperature of 30°C. Quartz surfaces have amorphous

cellulose layers on the top of them (Figure 46). After the addition of the flow of cysteine to

the cells, the frequency decreased slightly due to the fixation of cysteine on the amorphous

107

layer surface. Later, the frequency increased tremendously after the addition of the buffer.

This indicates the gradual detachment of the whole amorphous cellulose surface from the

surface of the Quartz. A further increase of the frequency was observed after the addition of

the CBHI enzyme (Figure 46)

Apparently, the layers were not stable enough for detecting the influence of the enzymes. One

possible reason for the almost complete detachment of the layer is that the –SH group of

cysteine reacts with gold of the quartz surfaces (underneath the regenerated amorphous layer)

and this leads to the removal of the amorphous layer on top.

Figure 46 The frequency and dissipation monitoring of the QCM-D regenerated

cellulose surface. The arrows indicate the injection of cysteine and CBHI, respectively;

and the asterisk the rinsing step with acetate buffer.

L-cysteine was replaced by ascorbate for two reasons: (i) to try to minimize the removal of

the amorphous cellulose layer and (ii) to try to reproduce the same conditions used in the

literature (Selig et al. 2015). The temperature used was 30°C. The injection of ascorbate alone

causes a tremendous increase in frequency and removal of the layer of amorphous cellulose.

The addition of CBH1 also caused a further increase in frequency. (Figure 47) A conclusion

to this experiment would be that ascorbate detaches the regenerated amorphous layer.

108

Figure 47 The frequency and dissipation monitoring of the QCM-D regenerated amorphous

cellulose surface.

109

V.3.4.Injecting LPMO enzymes and CBHI enzyme to the regenerated

amorphous cellulose surface

In our working conditions (Temperature 30°C at a flow of 0.1 ml/min), after the addition of

LPMO-FL enzyme (Figure 48) the frequency remained almost stable or decreased slightly,

which may indicate that the enzyme was adsorbed onto the cellulose surface, since the

frequency is linked to the mass of substrate present on the quartz surface. The rinsing step

after the LPMO injection resulted in an increase in frequency and a decrease in dissipation.

Thus, the increase of the frequency could indicate a decrease of the mass of cellulose present

on the quartz surface. After the addition of CBHI enzyme, the frequency continued

increasing, which again suggested the release of mass from the surface. Therefore, the LPMO

enzyme seems to act on the surface of the regenerated amorphous cellulose, attacking the

cellulose structure and causing the detachment of the amorphous cellulose from the surface of

the quartz.

Dissipation monitoring is linked to the viscosity of the material. It remained stable and then

decreased after the addition of the buffer indicating a decrease of the viscosity of the sample.

This fact could be explained as an increase in the swelling of the cellulose network by the

solvent, which confirmed the LPMO action.

Figure 48 The frequency and dissipation monitoring of the QCM-D regenerated

amorphous cellulose surface where LPMO-FL was injected.

110

V.4 Detecting the oxidative cleavage at the surface of cellulose using a

colorimetric method

The activities of LPMOs are relatively difficult to detect at the surface of cellulose. The

methods to measure the activities of LPMOs are costly, tedious or often reflect only an

apparent activity of the enzyme on the polysaccharide substrates. A simple method was

recently developed to measure the oxidative cleavage made by LPMOs directly at the surface

of the cellulose and chitin polymers (Wang et al. 2018). The method is based on quantifying

the ionic binding of cations to carboxyl groups formed by the action of type-1 (C1-oxidizing )

LPMOs. The method is based on the colorimetric detection and quantification of the

pyrocatechol violet (PV)-Ni2+

complex. Thus the Ni2+

ions will link to the carboxyl groups

resulting from the LPMO action. The complexation with the chromophore pyrocatechol violet

allowed to determine the (PV)-Ni2+

complex remaining in solution and, by comparison to the

starting (PV)-Ni2+

concentration (present in the control buffer), the Ni2+

bound to the

substrates can be determined.

V.4.1. Assessment of the method on Avicel and PASC

To validate the method, we tested the PaLPMO9E, which is an enzyme that oxidatively

cleave cellulose only at the C1-position (Bennati-Granier et al., 2015). The absorbance at 650

nm of the supernatant of the control containing only ethanol/HEPES was 0.28 (Figure 49).

This absorbance decreased in the supernatant of the reaction mixture consisting of PASC or

Avicel. This meant that the surface of PASC and Avicel interacted with the Ni2+

ions.

Interestingly, after adding PaLPMO9E and ascorbate to PASC, the absorbance further

decreased to reach a value of 0.14±0.025. This indicated that PaLPMO9E creates negative

charges via C1-oxidations at the surface of the PASC fibers which interact with Ni2+

ions and

decreases their presence in the surpernatant. In agreement with previous observations

(Bennati-Granier et al., 2015), we can observe using this colorimetric assay that PaLPMO9E

is less active on Avicel as compared to PASC.

111

Figure 49 Absorbance of the (PV)-Ni2+

complex present in the supernatant of different

samples treated or not with PaLPMO9E.

These experiments allowed concluding that this assay seems to be suitable to easily detect the

oxidations made by LPMOs at the surface of insoluble substrates.

V.4.2. Testing of LPMO-FL and LPMO-CD enzymes on PASC

The next step was to test the enzymes from our study (LPMO-FL and LPMO-CD), which

oxidatively cleave cellulose at both C1- and C4- positions. Since we observed no difference

between LPMO-FL and LPMO-CD on PASC with short incubation time (data not shown),

further experiments were carried out with longer incubation times (Figure 50).

After two days of incubation, we can observe a significant difference between the control

sample and the samples treated with either LPMO-CD of LPMO-FL. This effect seems to be

more pronounced in the presence of H2O2, which has been hypothesized to be the co-substrate

of LPMOs, therefore increasing the number of oxidative cleavage.

0

0.05

0.1

0.15

0.2

0.25

0.3A

bso

rban

ce a

t 6

50

nm

Samples

ControlEthanol/HEPES+NiCl2

PASC

PASC+LPMOE+Ascorbate

Avicel

Avicel+LPMOE+Ascorbate

112

Figure 50 Absorbance of (PV)-Ni2+

present in the supernatant of different samples with

LPMO-FL and LPMO-CD.

This determination confirmed the oxidation of cellulose upon the action of both LPMOs. The

colorimetric method is promising but we need to (i) repeat these preliminary results, (ii)

Quantify the number of oxidative cleavage (iii) test other LPMOs and (iv) test other substrates

(BMCC, NFC, Tunicates).

0

0.05

0.1

0.15

0.2

0.25

Samples after 16hours of

incubation

Samples after 2days of incubation

Ab

sorb

ance

at

65

0 n

m PASC

PASC+LPMO-FL+cysteine

PASC+LPMO-FL+H2O2

PASC+LPMO-CD+cysteine

PASC+LPMO-CD+H2O2

113

References

Dupont AL, and Mortha G. 2004. Comparative evaluation of size-exclusion chromatography

and viscometry for the characterisation of cellulose. Journal of Chromatography A

1026(1-2):129-141.

Henriksson M, Henriksson G, Berglund LA, and Lindstrom T. 2007. An environmentally

friendly method for enzyme-assisted preparation of microfibrillated cellulose (MFC)

nanofibers. European Polymer Journal 43(8):3434-3441.

Kohnke T, Ostlund Å, and Brelid H. 2011. Adsorption of arabinoxylan on cellulosic surfaces:

influence of degree of substitution and substitution pattern on adsorption

characteristics. Biomacromolecules 12(7):2633-2641.

Nechyporchuk O, Pignon F, and Belgacem MN. 2015. Morphological properties of

nanofibrillated cellulose produced using wet grinding as an ultimate fibrillation

process. Journal of Materials Science 50(2):531-541.

Paakko M, Ankerfors M, Kosonen H, Nykanen A, Ahola S, Osterberg M, Ruokolainen J,

Laine J, Larsson PT, Ikkala O et al. . 2007. Enzymatic hydrolysis combined with

mechanical shearing and high-pressure homogenization for nanoscale cellulose fibrils

and strong gels. Biomacromolecules 8(6):1934-1941.

Rodahl M, and Kasemo B. 1996. A simple setup to simultaneously measure the resonant

frequency and the absolute dissipation factor of a quartz crystal microbalance. Review

of Scientific Instruments 67(9):3238-3241.

Sauerbrey G. 1959a. Use of the vibrating quartz for thin film weighing and microweighing. Z

Phys 155:206.

Sauerbrey G. 1959b. Verwendung von Schwingquarzen zur Wägung dünner Schichten und

zur Mikrowägung. Zeitschrift für physik 155(2):206-222.

Selig MJ, Vuong TV, Gudmundsson M, Forsberg Z, Westereng B, Felby C, and Master ER.

2015. Modified cellobiohydrolase-cellulose interactions following treatment with lytic

polysaccharide monooxygenase CelS2 (ScLPMO10C) observed by QCM-D. Cellulose

22(4):2263-2270.

Villares A, Bizot H, Moreau C, Rolland-Sabaté A, and Cathala B. 2017a. Effect of xyloglucan

molar mass on its assembly onto the cellulose surface and its enzymatic susceptibility.

Carbohydrate polymers 157:1105-1112.

Villares A, Moreau C, Bennati-Granier C, Garajova S, Foucat L, Falourd X, Saake B, Berrin

J-G, and Cathala B. 2017b. Lytic polysaccharide monooxygenases disrupt the

cellulose fibers structure. Scientific Reports 7.

Villares A, Moreau C, Dammak A, Capron I, and Cathala B. 2015. Kinetic aspects of the

adsorption of xyloglucan onto cellulose nanocrystals. Soft Matter 11(32):6472-6481.

Villares A, Moreau Cl, Capron I, and Cathala B. 2013. Chitin nanocrystal-xyloglucan

multilayer thin films. Biomacromolecules 15(1):188-194.

Wang D, Li J, Wong AC, Aachmann FL, and Hsieh YS. 2018. A colorimetric assay to rapidly

determine the activities of lytic polysaccharide monooxygenases. Biotechnology for

biofuels 11(1):215.

Chapter VI

Discussion

117

VI.1. Importance of substrates to study LPMOs

The action of LPMO has generally been studied on partially solubilized cellulosic substrates

like PASC (Bennati-Granier et al. 2015) (Bey et al. 2013) (Fanuel et al. 2017) (Isaksen et al.

2014) (Karnaouri et al. 2017). But, we know the importance of the substrates to visualize the

action of the LPMO enzymes.

In this thesis, we aimed to study the effect of LPMO enzymes on other more relevant

cellulosic substrates showing key structural parameters to reveal the mechanism of action of

these enzymes. Hence, several cellulosic substrates were chosen to be representative of the

different types of cellulose found in nature. Indeed, due to its complexity, intact cellulose

found in nature is almost impossible to study with the techniques used nowadays. Several

cellulosic and nanocellulosic substrates that are derived from cellulose, which is treated

mechanically, enzymatically and/or chemically were considered in this project. BMCC

represents crystalline cellulose, PASC represents amorphous cellulose, NFC represents mixed

crystalline and amorphous regions. The closest to natural fibers are the kraft fibers but still

they are modified compared to natural cellulose. The importance of this spectrum of

substrates is that it represents many possible dispositions of the cellulosic fibers. This has a

great potential in revealing the effect of the LPMO enzymes.

In this thesis, we have shown that an LPMO harboring a CBM1 module seems to release more

soluble sugars on the amorphous-type substrate at 0.1% substrate concentration. When

removing the CBM1 module little or no effect was observed on the PASC substrate. This

indicates that the CBM1 module is essential to take the enzyme directly to the substrate, and

helps it to bind and target the amorphous regions. However, when increasing the substrate

concentration to 1%, LPMO-CD behaves similarly to LPMO-FL releasing significant

amounts of soluble sugars. This indicates that at low water content and higher substrate

concentration, the interaction between the enzymes without CBM and the substrate is higher

which reduces the need for CBM.

The range of cellulosic substrates used in this study also revealed that the established dogma

that LPMOs act on recalcitrant cellulose (crystalline regions) is only partly true. Indeed, we

clearly show that our AA9 LPMO can bind to crystalline cellulose when the enzyme harbors a

CBM1 but no cleavage seems to occur when tested in synergy with a CBH enzyme. These

results are in line with the observation made by Villares et al using the same AA9 LPMO but

differ from the observation made by Eibinger et al using AFM. In the latter, the authors used a

118

different LPMO from the AA9 family so it is possible that all LPMOs do not target the same

regions in cellulose. These results highlight the importance of the substrates chosen

representing crystalline or amorphous regions in order to compare the activities of LPMO-FL

and LPMO-CD. As a conclusion LPMO-FL binds to all types of substrates but acts mainly on

the amorphous regions of the insoluble substrates.

VI.2. Importance of experimental methods to assay LPMO activity

In the LPMO field, we are also aware of the importance of the methods to detect the effect of

the LPMO enzymes. Scientists generally use the Amplex red assay to determine the hydrogen

peroxide involved in the oxidative reaction, and high-performance anion exchange

chromatography (HPAEC) to identify and quantify the oxidized sugars released. The Amplex

red assay only gives information about the copper center and its reactivity, and it is usually

performed in the absence of substrate. On the other hand, HPAEC only allows measurement

of the soluble products released, which represent a very small percentage of the whole

cellulosic substrate. Therefore, there is a need for new methods to visualize the effect of the

LPMOs on cellulosic substrates.

In my thesis, I have used complementary techniques to the existing ones to visualize the effect

of LPMOs on the insoluble part of the cellulosic substrate remained after the enzymatic

action. The miscroscopy techniques, which were already set-up gave insights into the action

of the LPMO enzymes. On the other hand, I tried to determine the mass released after the

LPMO action by QCM-D but I experienced some difficulties with the QCM-D method

because of the instability of the regenerated cellulose layers and the non-reproducibility of the

frequency and dissipation graphs between the four cells of the QCM-D. Regarding the

HPSEC method, a major difficulty was the lack of reproducibility between the analyses of the

dissolved polymers. Indeed the dissolution process was not optimized and some parameters

(presence of water, time of agitation, accuracy of the HPSEC machine, data treatment and

integration of the peaks) influenced the results. Regarding the colorimetric assay used to

detect the oxidations at the surface of cellulose, results are promising but more experiments

would be needed to confirm the preliminary results obtained. An effort is needed to improve

these methods, but we are on the right path. As future prospects, additional methods could be

used: X-rays, solid state NMR, real-time AFM in liquid conditions, …

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VI.3. Importance of experimental conditions to assay LPMO activity

Another important consideration to take into account in the LPMO field is the set-up of

experimental conditions to analyze LPMO action. Indeed, the activity of LPMO enzymes is

highly influenced by the (i) substrate type, (ii) substrate concentration, (iii) electron donor

type, (iv) time of incubation, (v) the involvement of another enzyme like the CBHI, and (vi)

the addition of H2O2. Concerning the substrate type, LPMO-FL displayed highest activity on

the amorphous type substrate PASC and less activity was observed on the mixed-type

substrate NFC. As mentioned above, it seems that our LPMO enzyme acts on the amorphous

regions of cellulose, the most recalcitrant substrate remaining the crystalline-type substrate

BMCC. At first, when the CBM was deleted very little activity was obtained based on the

quantification of soluble products using HPAEC. Using another complementary enzyme,

CBHI, we revealed an improvement of its activity after the pretreatment with LPMO-CD.

This indicates the importance of the use of enzymes with complementary activities getting

inspired by co-secreted enzymes by filamentous fungi in nature. Changing the substrate

concentration revealed that the LPMO-CD displayed high activity (similar to LPMO-FL)

when we increased the concentration of the substrate to 1% (w/v). Concerning the electron

donor, ascorbate produced some peaks that interfered with the oxidized and non-oxidized

sugar peaks obtained by HPAEC. Cysteine was shown to be the most efficient electron donor

without producing any interfering peaks (Kracher et al. 2016). Finally, in the colorimetric

assay, H2O2 addition, revealed a decrease in the absorbance of Ni2+

of the supernatant of

LPMO-CD meaning that there is more Ni2+

ions fixed on the oxidations created on the surface

of the PASC substrate by LPMO-CD. Addition of H2O2 created even more oxidations and a

more important decrease of the absorbance. These preliminary results are in line with the

hypothesis drawn by (Bissaro et al. 2017) that LPMOs behave like peroxygenases using H2O2

as co-substrate.

All of these previously cited examples are evidence that the use of LPMO enzymes is highly

influenced by the experimental conditions. Therefore, a perspective of my work could be to

propose a more systematic way to study the influence of the experimental conditions to

investigate more deeply the action of LPMOs on cellulosic substrates.

VI.4. Depicting the impact of the CBM for LPMO action

The results obtained during my PhD demonstrate that the CBM contribute to the action of

LPMOs but it is not essential for enzyme activity. The functional relevance of CBMs and

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their contribution to the activity of LPMO enzymes had already been investigated (Crouch et

al. 2016);(Danneels et al. 2018), but in several cases, analysis surprisingly found modest and

contradictory effects on enzyme activity. The role of CBMs appended to glycoside hydrolases

has been explored in depth (Gilbert et al. 2013) and it is generally acknowledged that the

presence of CBMs increases the concentration of proteins at the surface of the substrate thus

increasing the activity of the enzyme (Guillen et al. 2010). In CAZymes, several enzymes

proved that the CBM takes the enzyme to the substrate.

In our work, the binding assay indicated that the enzyme with CBM binds to the all the

substrates tested. However, when the CBM was removed, the binding was not observed using

the pull-down assay technique, meaning that the binding was most probably weaker. This

means that the CBM drives the enzyme to the substrate in accordance with previous studies.

The binding of LPMO-FL with its CBM1 results in the release of soluble sugars products

even at low substrate concentration. However, the CBM seems to be beneficial for the

enzyme only at low substrate concentration.

VI.5. Which LPMOs for nanocelluloses production?

LPMOs were found to create defibrillation of kraft fibers (Villares et al., 2017). They have a

great potential in the production of nanocelluloses (Moreau et al. 2019). In order to select an

enzyme for large scale application experiments to produce nanocellulose at industrial scale,

one can wonder which LPMO would be ideal? Based on the results obtained in this thesis, it

is difficult to conclude on what is the most important criteria but definitely, we know that the

measurement of LPMO action on insoluble cellulose is the key parameter to reveal which

LPMO will be the most efficient for applied prospects. However, during this project, many

problems occurred when we tried to measure the impact of LPMOs on the insoluble fraction

of cellulose. In order to get more reliable data, AFM should be done on more samples maybe

50 samples for each enzyme in order to have statistically relevant results. This could give an

opportunity to reassure that LPMO-FL creates more tearing than LPMO-CD and that LPMO-

CD creates less tearings on the surface but interferes with the overall cohesion of the fiber’s

architecture. Liquid-phase atomic force microscopy is a very interesting technique to visualize

in real-time what the enzyme is causing on the surface of the fibers. The time-course variation

of the fiber’s aspects would be a clear proof that the tearings are caused by the enzyme and

not by earlier pretreatments. If tearings caused by oxidation are observed more frequently on

the LPMO-FL-treated sample then LPMO-FL should be the chosen most efficient enzyme for

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larger scale applications. Although there should be a restriction concerning the size of the

fibers tested so not all the substrates are adequate for the technique. That’s why optimizing

the solubilization of the substrates in DMAc/LiCl could be a good alternative for larger fibers.

Figuring out the effect of the enzymes on the weight-average molecular weight of the fibers is

an excellent approach to choose the best enzyme. The key parameter is to solubilize the fibers.

There must be an improvement of the solubilization of the fibers in DMAc/LiCl and a

repetition of the samples for several times to have statistically relevant results. Then, the

enzymes having the highest impact on the DP of the fibers would be chosen and could be

tested at higher scale.

122

References

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D, Rogniaux H, Gimbert I et al. . 2015. Substrate specificity and regioselectivity of

fungal AA9 lytic polysaccharide monooxygenases secreted by Podospora anserina.

Biotechnology for Biofuels 8.

Bey M, Zhou S, Poidevin L, Henrissat B, Coutinho PM, Berrin J-G, and Sigoillot J-C. 2013.

Cello-Oligosaccharide Oxidation Reveals Differences between Two Lytic

Polysaccharide Monooxygenases (Family GH61) from Podospora anserina. Applied

and Environmental Microbiology 79(2):488-496.

Bissaro B, Rohr AK, Muller G, Chylenski P, Skaugen M, Forsberg Z, Horn SJ, Vaaje-Kolstad

G, and Eijsink VGH. 2017. Oxidative cleavage of polysaccharides by monocopper

enzymes depends on H2O2. Nature Chemical Biology 13(10):1123-+.

Crouch LI, Labourel A, Walton PH, Davies GJ, and Gilbert HJ. 2016. The Contribution of

Non-catalytic Carbohydrate Binding Modules to the Activity of Lytic Polysaccharide

Monooxygenases. Journal of Biological Chemistry 291(14):7439-+.

Danneels B, Tanghe M, and Desmet T. 2018. Structural Features on the Substrate-Binding

Surface of Fungal Lytic Polysaccharide Monooxygenases Determine Their Oxidative

Regioselectivity. Biotechnology journal.

Fanuel M, Garajova S, Ropartz D, McGregor N, Brumer H, Rogniaux H, and Berrin J-G.

2017. The Podospora anserina lytic polysaccharide monooxygenase PaLPMO9H

catalyzes oxidative cleavage of diverse plant cell wall matrix glycans. Biotechnology

for Biofuels 10.

Gilbert HJ, Knox JP, and Boraston AB. 2013. Advances in understanding the molecular basis

of plant cell wall polysaccharide recognition by carbohydrate-binding modules.

Current Opinion in Structural Biology 23(5):669-677.

Guillen D, Sanchez S, and Rodriguez-Sanoja R. 2010. Carbohydrate-binding domains:

multiplicity of biological roles. Applied Microbiology and Biotechnology 85(5):1241-

1249.

Isaksen T, Westereng B, Aachmann FL, Agger JW, Kracher D, Kittl R, Ludwig R, Haltrich

D, Eijsink VGH, and Horn SJ. 2014. A C4-oxidizing Lytic Polysaccharide

Monooxygenase Cleaving Both Cellulose and Cello-oligosaccharides. Journal of

Biological Chemistry 289(5):2632-2642.

Karnaouri A, Muraleedharan MN, Dimarogona M, Topakas E, Rova U, Sandgren M, and

Christakopoulos P. 2017. Recombinant expression of thermostable processive Mt EG5

endoglucanase and its synergism with Mt LPMO from Myceliophthora thermophila

during the hydrolysis of lignocellulosic substrates. Biotechnology for biofuels

10(1):126.

Kracher D, Scheiblbrandner S, Felice AKG, Breslmayr E, Preims M, Ludwicka K, Haltrich

D, Eijsink VGH, and Ludwig R. 2016. Extracellular electron transfer systems fuel

cellulose oxidative degradation. Science 352(6289):1098-1101.

Moreau C, Tapin-Lingua S, Grisel S, Gimbert I, Le Gall S, Meyer V, Petit-Conil M, Berrin J-

G, Cathala B, and Villares A. 2019. Lytic polysaccharide monooxygenases (LPMOs)

facilitate cellulose nanofibrils production. Biotechnology for Biofuels 12.

123

Annexes

Supplementary figure S1: optimized gene sequence for the expression in Pichia pastoris

encoding for the PaLPMO9H.

Supplementary figure S2: AFM topographical 3D images of the control kraft fibers (a-

b), of LPMO-FL-treated fibers (c-d) and of LPMO-CD-treated fibers (e-f) without

mechanical dispersion.

Résumé de la thèse en français

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Ce projet de thèse porte sur les « lytic polysaccharide monooxygenases » (ou LPMO) qui sont

des enzymes fongiques récemment découvertes agissant sur le polymère le plus abondant sur

terre : la cellulose. Les objectifs de cette thèse sont de comprendre le mécanisme d’action des

enzymes LPMO sur les substrats lignocellulosiques : cellulose microcristalline bactérienne,

nanofibrilles de cellulose, cellulose amorphe et cellulose kraft. Les modifications effectuées

par ces enzymes seront étudiées à différentes échelles : micrométrique, moléculaire et

nanométrique.

I. Introduction

I.1 Structure moléculaire de la cellulose

Figure 1: Structure moléculaire de la cellulose (n=DP, degré de polymérisation).

Depuis sa découverte par Payen, les caractéristiques physiques et chimiques de la cellulose

ont été intensivement étudiées. La cellulose est un homopolymère glucidique formé d'unités

répétitives de molécules de β-D-glucopyranose liées de manière covalente par des liaisons

glycosidiques entre le groupe OH équatorial de l'atome de carbone C4 et l'atome de carbone

C1 (β-1,4-glucane) (Figure 1).

L'unité chimique répétitive est donc la molécule de D-glucopyranose et l'unité structurale

répétitive est le cellobiose, formé de deux monosaccharides successifs. (Lavoine et al. 2012b);

(Klemm et al. 2005).

I.2. Organisation hiérarchique de la cellulose

Les trois groupes hydroxyles (groupes OH) de l'unité de glucose peuvent former des liaisons

hydrogène intra et inter moléculaires. La force de ces liaisons hydrogène est d’environ 25

kJ/mol. Elles sont responsables de la rigidité de la molécule de cellulose (Habibi et al. 2010).

Les fibrilles élémentaires sont regroupées dans des unités plus grandes appelées microfibrilles

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de cellulose. Les microfibrilles de cellulose sont assemblées en fibres de cellulose qui sont

représentées dans la figure 2. Le fractionnement à l’échelle nanométrique permet la

production des nanocelluloses. Il existe deux types principaux de nanocelluloses: (i) les

nanocristaux de cellulose et (ii) les microfibrilles de cellulose (Lavoine et al. 2012a)

Figure 2: La cellulose : des sources jusqu’aux molécules: (Lavoine et al. 2012b).

I.3 Les substrats cellulosiques

I.3.1 Phosphoric acid swollen cellulose (PASC)

Le gonflement de la cellulose dans de l'acide orthophosphorique permet la fabrication de la

« phosphoric acid swollen cellulose » ou PASC qui du fait de son accessibilité est un substrat

très utilisée pour les études de dégradation de la cellulose. La PASC est constituée de chaine

de cellulose amorphes. Le gonflement dans l'acide phosphorique perturbe les liaisons

hydrogène ordonnées dans la cellulose cristalline, ce qui entraîne une amorphisation des

chaînes.

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I.3.2 Nanocristaux de cellulose bactérienne (NCB) et nanocristaux de cellulose

(NCC)

La cellulose bactérienne est obtenue à partir d'espèces appartenant aux genres Acetobacter,

Rhizobium, Agrobacterium et Sarcina par le biais de diverses méthodes et techniques de

culture, l'Acetobacter xylinum étant la plus étudiée.

Les nanocristaux de cellulose (Figure 3) ont été produits au début par Bengt G. Ranby (Rånby

1951) en traitant de la pâte de sulfite blanchie et raffinée avec de l’acide sulfurique.

Lorsqu'une combinaison appropriée de la concentration en acide, du temps et de la

température est utilisée, les régions amorphes de la fibre de cellulose sont hydrolysées plus

rapidement que les régions cristallines, ce qui permet d'isoler les nanocristaux de cellulose

(Habibi et al. 2010).

Figure 3: Images TEM de la dispersion séchée de nanocristaux de cellulose provenant de

(a) tunicier, (b) cellulose bactérienne, (c) ramie, (d) sisal (Habibi et al. 2010).

Les NCB présentent un rapport de forme élevé avec des sections transversales rectangulaires

ou une forme en ruban. L'utilisation d'acide chlorhydrique produit des nanocristaux neutres

qui ne sont pas stabilisés par répulsions électrostatiques. (Pirich et al. 2015), Les NCB

semblent être groupés en fagots ou agrégés.

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I.3.3 Nanofibrilles de cellulose (NFC)

Les NFC ont d'abord été produits par Turbak en 1983 par délamination mécanique des fibres

de cellulose (Turbak et al. 1983). La fibrillation libère de longues fibrilles flexibles

constituées de régions alternées cristallines et amorphes, d’une largeur d’environ 5-20 nm et

d’une longueur de plusieurs microns. Les NFC présentent un rapport de forme élevé et forme

des réseaux enchevêtrés pouvant former des gels.

Actuellement, il existe un grand nombre de procédés de désintégration mécanique différents

pour produire du NFC. Ils peuvent être subdivisés en procédés conventionnels

(homogénéisation, microfluidisation et raffinage) et non conventionnels (extrusion, explosion

à la vapeur, ultrasonication broyage à billes, etc.). L'homogénéisation à haute pression est la

plus couramment utilisée. Elle consiste à faire passer la suspension de cellulose à travers un

espace réduit entre la vanne d'homogénéisation et un anneau de choc (Nechyporchuk et al.

2016).

Les principaux obstacles à la production de NFC sont la consommation élevée d’énergie et le

blocage de l’homogénéisateur par des particules de taille supérieures à l’espace disponible

dans la vanne. (Lindstrom 2017).

Afin de réduire l’énergie impliquée dans le délaminage mécanique des fibres de cellulose,

différents prétraitements enzymatique et chimique sont utilisés dans l’industrie. (Saito and

Isogai 2004)

I.3.4 Fibres de pâte kraft

Dans l’industrie papetière, la fabrication de la pâte consiste à séparer les fibres de cellulose

des autres composants de la paroi cellulaire en ayant le plus faible impact sur la fibre. Cette

séparation peut être réalisée par des traitements mécaniques ou par des réactions chimiques,

ainsi que par une combinaison des deux. La pâte kraft est la forme la plus courante de pâte

chimique, représentant 80% de l’ensemble de l’industrie de la pâte chimique (Saito and Isogai

2004) (Chevalier-Billosta 2008)

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I.4 Dégradation enzymatique fongique de la cellulose.

La plupart des microorganismes cellulolytiques sécrètent une grande variété d’enzymes

lorsqu’ils sont cultivés sur des substrats cellulosiques, parmi lesquelles figurent les enzymes

hydrolitiques et les enzymes ayant des activités auxiliaires (oxydases) qui agissent en synergie

avec les cellulases (voir Figure 4).Récemment, des enzymes oxydatives appelées LPMO («

lytic polysaccharide monooxygenase ») ont révolutionnées notre vision de la dégradation de

la cellulose. Suite au clivage oxydatif à la surface de la cellulose, les LPMO sont capables de

défibriller la cellulose, fragilisant ainsi sa structure et facilitant l’action des enzymes

hydrolytiques. A ce jour, les LPMO sont classés dans 7 familles AA de la classification CAZy

(www.cazy.org) sachant que la famille AA9 est retrouvée uniquement chez les champignons

et que les membres de cette famille sont tous actifs sur la cellulose.

Figure 4 Schéma actuel de la dégradation enzymatique fongique de la cellulose. A noter

que de nombreux systèmes enzymatiques cellulolytiques ont de multiples

Endoglucanases (EG) et / ou cellobiohydrolases (CBH) pouvant agir sur différentes

parties du substrat avec des différences en terme de crystalinité et d’accessibilité. La

cellobiose déshydrogénase (CDH) peut fournir des électrons aux LPMO AA9. Il a été

également démontré que d’autres réducteurs non enzymatiques (donneurs d’électrons)

induisaient une activité oxydative (par exemple, acide ascorbique, lignine) (Horn et al.

2012).

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II. matériel et méthodes

II.1. Substrats cellulosiques

Plusieurs substrats cellulosiques ont été utilisés dans cette étude. Ils représentent soit les

régions cristallines (NCB), amorphes (PASC), les régions alternées cristallines et amorphes.

(NFC obtenues via un prétraitement par endoglucanase suivi d'une microfluidisation), ou très

proches des fibres naturelles comme le papier kraft délignifié d'arbres résineux. Des

cellooligosaccharides ont été également utilisés comme substrats.

II.2. Production hétérologue d'enzymes chez Pichia pastoris

II.2.1. Souches et milieux :

La souche X33 de P. pastoris et le vecteur pPICZαA sont utilisés dans cette étude. La souche

SuperMan 5 de Pichia pastoris produit des protéines recombinantes avec moins de

glycosylation (Pekarsky et al. 2018).

II.2.2. Gènes synthétiques et vecteurs utilisés

Les gènes codant pour PaLPMO9H sont optimisés par l’usage des codons pour leur

expression dans Pichia pastoris. Ils ont été synthétisés par Genewiz (South Plainfield USA) et

insérés dans le vecteur pPICZαA en utilisant les sites de restriction BstBI et Xbal.

II.2.3.Transformation de Pichia pastoris.

Les cellules compétentes de P. pastoris ont été décongelées sur de la glace et transformées par

0,5-5 µg du plasmide linéarisé. Une tension de 1500 V a été appliquée sur de la glace pendant

5 ms à l'aide d'un électroporateur à microspulseurs (Biorad Marned-la-Coquette, France).

II.2.4.Production recombinante d'enzymes recombinantes

PaLPMO9H (protéine ID CAP 61476) et une enzyme modifiée sans CBM ont été produites

dans Pichia pastoris en utilisant des flacons (Bennati-Granier et al. 2015). Le milieu YPD

inoculé avec une oese de cellules de Pichia de la boîte de Pétri sous agitation à 160 tr / min à

30 ° C pendant quatre heures. Le milieu de production des protéines est le BMGY à 30 ° C et

à 200 tr / min. Après centrifugation, le culot cellulaire a été remis en suspension dans 1/5 de

milieu BMMY contenant du méthanol. L'induction de la production de l'enzyme a été réalisée

à 30 ° C pendant trois jours sous agitation constante à 200 tr / min et par addition de 3% de

méthanol chaque jour pendant trois jours. Chaque enzyme recombinante marquée (His)6 a été

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purifiée sur une colonne HiTrap HP et dialysées contre un tampon d’acétate de sodium 50

mM, pH 5,2.

II.3. Analyse des protéines

Les concentrations en protéines ont été déterminées par absorption à 280 nm en utilisant un

spectrophotomètre Nanodrop ND-2000 (Thermo Fisher Scientific). L'électrophorèse de

protéines a été réalisée en chargeant les protéines sur des gels SDS-PAGE préfabriqués

(Biorad, Marnes-la Coquette, France) et colorées au bleu de Coomassie. L'analyse ICP-MS a

été effectuée comme décrit dans (Couturier et al. 2018) pour quantifier la teneur en cuivre de

la protéine. Une analyse fluorimétrique à base de « Amplex red » et de « horseradish

peroxidase » a été utilisée comme décrit précédemment (Isaksen et al. 2014; Kittl et al. 2012).

II.4. Tests de liaison à la cellulose

Les mélanges réactionnels ont été réalisés avec une charge de substrat insoluble de 0,3% (p /

v) (BMCC; NFC; PASC) et 30 ug de protéines ont été ajoutés. Après centrifugation à 14 000

g pendant 10 min, le surnageant (contenant les protéines non liées) a été soigneusement

éliminé. Ensuite, les culots de polysaccharide ont été lavées deux fois (lavage 1 et lavage 2)

en les remettant en suspension dans du tampon et centrifugées à 14 000 g pendant 10 min.

Cette étape a été répétée deux fois. Le culot restant a finalement été remis en suspension dans

du tampon de charge SDS sans colorant et bouilli pendant 10 minutes pour dissocier toute

protéine liée. Les fractions non liées, de lavage 2 et liées ont été analysées par SDS-PAGE

pour détecter la présence ou l'absence de la protéine.

II.5. Essais enzymatiques

Tous les tests de clivage contenaient 0,1% (p/v) de substrat (PASC, BMCC, NFC) 4,4 µM de

PaLPMO9 et 1 mM de L-cystéine dans du tampon acétate de sodium 50 mM pH 5,2. Les

réactions enzymatiques ont été incubées dans un « thermomixer » (Eppendorf, Montesson,

France) à 50 ° C et 850 tr / min pendant 16 heures. À la fin de la réaction, les échantillons ont

été bouillis à 100 ° C pendant 15 min, puis centrifugés à 15 000 g pendant 10 min pour

séparer les fractions solubles et insolubles. Des analyses à une concentration de PASC de 1%

(p / v) ont également été effectuées avec les conditions mentionnées précédemment.

II.5.1. Essais combinés

Les analyses enzymatiques des LPMO ont été effectuées séquentiellement avec une

cellobiohydrolase de T. reesei (CBH-I) comme décrit dans (Filiatrault-Chastel et al. 2019).

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II.5.2. Analyse des oligosaccharides

Les cello-oligosaccharides oxydés et non oxydés générés après l'action de la LPMO ont été

analysés par chromatographie à haute performance d'échange d'anions couplée à une détection

ampérométrique pulsée (HPAEC-PAD) (ThermoFischer Scientific, États-Unis) (Bennati-

Granier et al. 2015).

II.6. Analyse de la fraction insoluble

II.6.1. Microscopie

Les échantillons de contrôle et les échantillons traités aux LPMO ont été analysés par

microscopie optique, microscopie à force atomique (AFM) et microscopie électronique à

transmission (MET). Une analyse plus approfondie a été réalisée par résonance magnétique

nucléaire (RMN), spectroscopie infrarouge à transformée de Fourier (IRTF) et une analyse

colorimétrique dont l'objectif principal est la détermination de la liaison des cations (Ni2+

) aux

groupes carboxyle formés par l'action des LPMO oxydants C1 sur les polysaccharides (Wang

et al. 2018)).

II.6.2. Chromatographie d'exclusion stérique haute performance couplée à un

détecteur de diffusion de lumière multi-angle (HPSEC-MALLS)

Les fibres kraft ont été incubées avec les enzymes à 50 ° C sous agitation douce pendant 16 h.

Plus tard, les échantillons ont été bouillis pendant 15 minutes. La moitié des échantillons a été

dispersée par un homogénéisateur Polytron et ultérieurement par ultrasons. Les fibres ont été

dissoutes par échange de solvant dans de l'eau, puis par dispersion dans du méthanol anhydre

(50 ml à chaque fois, suivies de trois redispersions supplémentaires dans du

diméthylacétamide anhydre (50 ml)). Ensuite, les fibres ont été ajoutées à 5 ou 10 ml de

DMAc. / LiCl (9% p / p) sous agitation mécanique pendant 24 heures avant dilution 10 fois

avec du DMAc anhydre (Dupont and Mortha 2004). Les solutions ont ensuite été filtrées et

injectées sur un système de chromatographie par exclusion stérique (OMNISEC Resolve,

Malvern) avec N , N-diméthylacétamide / chlorure de lithium (0,9% p / v) comme éluant pour

effectuer une analyse chromatographique couplée à un détecteur de diffusion de lumière et un

détecteur réfractométrique.

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III. Resultats

III .1 Production de PaLPMO9H avec et sans CBM1

Pour mieux comprendre la contribution du CBM1 à la fonction catalytique des LPMO AA9,

nous avons sélectionné PaLPMO9H sur la base des analyses biochimiques précédentes

(Bennati-Granier et al. 2015) (Villares et al. 2017) (Fanuel et al. 2017). PaLPMO9H est une

enzyme modulaire à deux domaines contenant un domaine AA9 catalytique N-terminal (16-

243) et un domaine CBM1 C-terminal (271-307) (Figure 5). Ces deux domaines sont liés via

un « linker » riche en sérine / thréonine / asparagine comprenant 27 résidus d'acides aminés.

Lorsque l'enzyme PaLPMO9H a été tronquée juste après le module catalytique en position

244, nous avons été incapables de produire avec succès la protéine recombinante chez P.

pastoris (données non présentées). Par conséquent, nous avons décidé de laisser 16 résidus

d’acides aminés du « linker » pour promouvoir la production de l’enzyme recombinante. En

utilisant cette stratégie, nous avons réussi à produire l'enzyme PaLPMO9H sans CBM1

tronquée en position 259. Dans la suite de l’étude, PaLPMO9H avec le CBM1 est nommé

LPMO-FL (entière), et le PaLPMO9H sans le CBM1 s'appelle LPMO-CD (domaine

catalytique). Comme prévu, la suppression du CBM1 a diminué la masse moléculaire de

l’enzyme de ~ 38 kDa (LPMO-FL) a ~ 33 kDa (LPMO-CD).

Figure 5. Representation schématique des enzymes utilisées. LPMO-FL (entière) et

LPMO-CD (domaine catalytique) tronquée au niveau du “linker” avec numérotation

des acides aminés des limites de chaque domaine.

La masse moléculaire apparente était encore légèrement supérieure à la masse moléculaire

théorique (25,7 kDa) en raison des glycosylations de O et N-. LPMO sont des enzymes

dépendantes du cuivre, ce qui rend crucial de vérifier le chargement correct en protéines de

cuivre. La charge en cuivre dans chaque enzyme a été quantifiée en utilisant La spectrométrie

136

de masse à plasma à couplage inductif, ou (ICP-MS). Tous les deux les enzymes étaient

également chargées d'environ ~ 1 atome de cuivre par molécule (à savoir 10,3 et 10,8 µM de

Cu2 +

pour 10 µM de LPMO-FL et LPMO-CD, respectivement).

III.2. L’absence de CBM1 altère l’activité cellulolytique de la LPMO à faible

concentration en substrat.

L’action de LPMO-FL a été évaluée au début sur trois types de cellulose : cellulose gonflée à

l'acide phosphorique « PASC », cellulose nanofibrillée « NFC », et cellulose microcristalline

bactérienne « BMCC » (Figure 6a). Comme indiqué précédemment, LPMO-FL a libéré à la

fois des molécules oxydées en C4 (C4ox) et des oligosaccharides non oxydés de la PASC

[18]. Cependant, l’utilisation de NFC comme substrat a conduit à moins de produits libérés, et

l’utilisation d’un substrat cristallin plus récalcitrant (BMCC) conduit à des produits à peine

détectables (Figure 6a). Nous avons ensuite comparé l’action des deux LPMO-FL LPMO-CD

en mesurant la libération de sucres provenant de PASC (Figure. 6b). LPMO-FL a libéré des

quantités plus élevées de sucres solubles (oligosaccharides oxydés et non oxydés) comparé au

LPMO-CD où les sucres solubles étaient à peine détectables (Figure 6b). LPMO-FL étant

actif sur les oligosaccharides solubles (Bennati-Granier et al. 2015), nous avons étudié

l’activité de LPMO-FL et de LPMO-CD sur le cellohexaose en tant que substrat. Une analyse

au cours de temps a révélé que les deux enzymes ont été capables de cliver le cellohexaose,

conduisant principalement aux produits non oxydés Glc3 et Glc4 et Produits oxydés en C4.

Bien que LPMO-FL ait montré activité légèrement supérieure à celle du LPMO-CD en 24 h

dans le temps, le clivage observé du cellohexaose par LPMO-CD confirme que l’enzyme

dépourvue du module CBM1 est toujours fonctionnelle.

La liaison LPMO-FL et LPMO-CD à PASC, BMCC et NFC a été évaluée en l’absence de

réducteur en utilisant le test « pull down » pour évaluer l'impact du CBM1. LPMO-FL a été

observée dans la fraction liée des trois substrats cellulosiques testés. Cependant, en l’absence

de CBM1, il n’existait pas bandes correspondantes à LPMO-CD dans la fraction liée. Par

conséquent, le CBM1 favorise la liaison entre LPMO et les substrats cellulosiques.

137

Figure 6: Analysis des produits de degradation solubles.a Produits libérés par LPMO-

FL à partir de PASC, NFC ou BMCC. b Analyse des produits de degradation solubles

libérés par LPMO-FL et LPMO-CD à partir de PASC.

10 15 20 25 30 35 40 45

0

200

400

600

800

Charg

e [

nC

]

Retention Time [min]

PASC

NFC

BMCC

PASC (no enzyme)

C4oxC4ox C1-C4ox

Glc

3

Glc

6

Glc

4

Glc

5

Glc

2

Glc2-Glc6

10 15 20 25 30 35 40 45

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800

Charg

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nC

]

Retention Time [min]

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LPMO-CD

No enzyme

200

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600

Glc

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Glc

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Glc

6

Glc

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Glc

5

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a

b

138

III.3.Action combinée de LPMO-FL et LPMO-CD avec une cellobiohydrolase

Pour évaluer l'impact du LPMO-CD sur les substrats cellulosiques, nous avons testé les

enzymes LPMO-FL et LPMO-CD en association avec la cellobiohydrolase qui agit sur la

terminaison réductrice. (famille GH7 CBH-I) de T. reesei. Les substrats cellulosiques ont été

prétraités séquentiellement avec LPMO-FL ou LPMO-CD avant l’ajout de l’enzyme CBH-I

Étant donné que les LPMO et CBH-I agissent sur des substrats solubles, nous avons mis en

place une étape de lavage après le traitement LPMO pour évaluer l’impact du traitement

LPMO uniquement sur les fibres insolubles. Le prétraitement au LPMO était bénéfique sur le

PASC et le NFC mais n’a eu aucun effet visible sur le substrat cristallin BMCC (Figure 7). Le

prétraitement avec soit LPMO-CD ou LPMO-FL entraine une augmentation de la libération

de cellobiose d'environ 30% du substrat NFC. Cependant, le prétraitement au LPMO-FL était

plus efficace sur la PASC (augmentation de 60% de la production de cellobiose) par rapport à

la LPMO-CD. Pris ensemble, ces résultats montrent qu’aucune des deux LPMO ne cible la

fraction cristalline de la cellulose. Nous pensons que les deux LPMO ciblent des régions

amorphes facilitant ainsi l'activité de CBH-I sur la cellulose crystalline. De plus, dans le cadre

de ces expériences, la présence du module CBM1 n'est pas strictement nécessaire pour l'action

de la LPMO.

Figure 7: Action combinée de LPMO-FL et LPMO-CD avec la cellobiohydrolase (CBH)

pour la degradation de NFC, PASC and BMCC.

139

III.4 L'augmentation de la concentration en substrat impacte l’effet du CBM1

L’étape suivante consistait à déterminer si la concentration de substrat a une influence sur

l'activité des enzymes. Nous avons augmenté la concentration du substrat à 1% (p / v) pour

favoriser la probabilité d'interactions enzyme-substrat dans un contexte sans CBM. À forte

concentration en substrat, les sucres solubles libérés par LPMO-CD sont devenus détectables

et étaient dans la même gamme que les sucres solubles libérés par LPMO-FL à partir de la

PASC (Figure 8). Fait intéressant, les produits oxydés en C1 (C1ox) (Glc2ox-Glc4ox), qui

étaient à peine détectables avec LPMO-FL, ont été abondamment libérés par LPMO-CD

(Figure 8). Les produits oxydés en C4 élués à environ 30 min étaient moins abondants alors

que les produits oxydés en C1 / C4 éluant entre 41 et 42 min ont légèrement augmenté.

L'absence de CBM induit donc une modification de régiosélectivité de l’enzyme (Figure 8).

Figure 8: Analyse par chromatographie ionique HPAEC des produits de degradation

libérés par les enzymes LPMO-FL and LPMO-CD.

140

III.5. Impact du CBM sur la fraction insoluble

Pour tenter de mieux comprendre le rôle de du CBM sur l’action des LPMO, nous avons

évalué les changements de morphologie de fibres kraft dus à l'incubation avec les LPMO.

Nous avons d’abord étudié la structure de la fibre en utilisant la microscopie optique. Les

fibres kraft d'origine mesurent des dizaines de micromètres de diamètre et environ 1 mm de

long (Figure. 9a). Après traitement LPMO, il n'y avait pas de changements visibles dans

l’aspect physique des fibres, c'est-à-dire morphologie fibreuse ou dimensions, dans les

échantillons traités LPMO-FL (Figure. 9b) et LPMO-CD. Comme décrit précédemment

(Villares et al. 2017), l’action des LPMOs ne produit pas à elle seule une différence notable

dans la désintégration des fibres kraft, de la même manière que l'action de cellulases (Paakko

et al. 2007) (Henriksson et al. 2007) (Nechyporchuk et al. 2015). Par conséquent, après le

traitement LPMO, les fibres ont été dispersées mécaniquement puis soumises à un traitement

court par ultrasons. La dispersion a révélé l’effet du LPMO sur les fibres kraft. Les

échantillons de contrôle ont montré légère défibrillation alors que les deux échantillons traités

au LPMO ont montré une délamination claire de la paroi cellulaire (Figure. 9d – f). LPMO-FL

et LPMO-CD semblaient tous deux créer des points faibles dans la fibre qui ont facilité la

désintégration mécanique. Pour obtenir une meilleure image de l’effet de la LPMO, nous

avons utilisé la microscopie à force atomique (AFM) pour analyser les échantillons (Figure.

9g – i). Les images topographiques ont montré la présence de grosses fibres dans les

échantillons de contrôle et dissociation nette dans des échantillons traités aux LPMO. LPMO-

FL induit la fibrillation des fibres kraft qui forment un réseau enchevêtré de nanofibrilles

d’environ 5 nm de diamètre. LPMO-CD a également produit un réseau de fibres désintégrées

mais avec des faisceaux de fibres plus épais. En comparant l'apparence des fibres traitées avec

LPMO-FL ou LPMO-CD contre les contrôles, il est évident que les deux enzymes influencent

la cohésion et l’architecture des fibres, ce qui les rend plus sensibles aux forces mécaniques

causées par la dispersion. En se basant sur les images AFM, les deux LPMO ont réduit la

cohésion de la fibre, mais la présence de CBM apparemment permis à LPMO-FL de

défibriller la cellulose.

141

Figure. 9 Morphologie des fibres kraft traitées aux LPMOs. Images de microscopie

optique de fibres kraft avant (a-c) et après (d-f) dispersion mécanique pour les

échantillons témoins (a,d), traités avec LPMO-FL (b,e) et les fibres traitées avec LPMO-

CD (c,f). Images topographiques AFM après traitement LPMO et dispersion pour les

fibres kraft témoins (g), les fibres traitées aux LPMO-FL (h) et les fibres traitées au

LPMO-CD (i).

142

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Chalak et al. Biotechnol Biofuels (2019) 12:206 https://doi.org/10.1186/s13068-019-1548-y

RESEARCH

Influence of the carbohydrate-binding module on the activity of a fungal AA9 lytic polysaccharide monooxygenase on cellulosic substratesAmani Chalak1,2, Ana Villares1, Celine Moreau1, Mireille Haon2, Sacha Grisel2, Angélina d’Orlando1, Isabelle Herpoël‑Gimbert2, Aurore Labourel2, Bernard Cathala1 and Jean‑Guy Berrin2*

Abstract

Background: Cellulose‑active lytic polysaccharide monooxygenases (LPMOs) secreted by filamentous fungi play a key role in the degradation of recalcitrant lignocellulosic biomass. They can occur as multidomain proteins fused to a carbohydrate‑binding module (CBM). From a biotech perspective, LPMOs are promising innovative tools for produc‑ing nanocelluloses and biofuels, but their direct action on cellulosic substrates is not fully understood.

Results: In this study, we probed the role of the CBM from family 1 (CBM1) appended to the LPMO9H from Podos-pora anserina (PaLPMO9H) using model cellulosic substrates. Deletion of the CBM1 weakened the binding to cellulose nanofibrils, amorphous and crystalline cellulose. Although the release of soluble sugars from cellulose was drastically reduced under standard conditions, the truncated LPMO retained some activity on soluble oligosaccharides. The cel‑lulolytic action of the truncated LPMO was demonstrated using synergy experiments with a cellobiohydrolase (CBH). The truncated LPMO was still able to improve the efficiency of the CBH on cellulose nanofibrils in the same range as the full‑length LPMO. Increasing the substrate concentration enhanced the performance of PaLPMO9H without CBM in terms of product release. Interestingly, removing the CBM also altered the regioselectivity of PaLPMO9H, signifi‑cantly increasing cleavage at the C1 position. Analysis of the insoluble fraction of cellulosic substrates evaluated by optical and atomic force microscopy confirmed that the CBM1 module was not strictly required to promote disrup‑tion of the cellulose network.

Conclusions: Absence of the CBM1 does not preclude the activity of the LPMO on cellulose but its presence has an important role in driving the enzyme to the substrate and releasing more soluble sugars (both oxidized and non‑oxidized), thus facilitating the detection of LPMO activity at low substrate concentration. These results provide insights into the mechanism of action of fungal LPMOs on cellulose to produce nanocelluloses and biofuels.

Keywords: Filamentous fungi, Cellulose, LPMO, CBH, CBM, Microscopy

© The Author(s) 2019. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creat iveco mmons .org/licen ses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Open Access

Biotechnology for Biofuels

*Correspondence: jean‑guy.berrin@inra.fr2 Biodiversité et Biotechnologie Fongiques, UMR1163, INRA, Aix Marseille Université, Marseille, FranceFull list of author information is available at the end of the article

Page 2 of 10Chalak et al. Biotechnol Biofuels (2019) 12:206

BackgroundCellulose is the most abundant biopolymer on Earth and one of the main sources of renewable carbon [1]. Huge effort is being invested in the development of biofuels made from cellulosic biomass feedstocks, known as sec-ond-generation biofuels [2]. In parallel, nanomaterials such as nanofibers and nanocrystals are being isolated from wood and agricultural resources by mechanical and/or chemical treatments, offering unique properties with a wide range of applications (paper, pharmaceutical, cosmetics and food industries) [3–5]. The hierarchical complexity and recalcitrance of cellulose create a need to process it via innovative “green” pretreatments to address pressing global challenges and environmental concerns.

In nature, cellulose degradation is mainly achieved by filamentous fungi, which secrete complementary hydro-lytic and oxidative activities. In contrast to known cellu-lases, which are hydrolytic enzymes, lytic polysaccharide monooxygenases (LPMOs) degrade cellulose via an oxi-dative mechanism [6–8] involving molecular oxygen or hydrogen peroxide and redox-active molecules acting as electron donors [9, 10]. LPMO-catalyzed cleavage leads to oxidation of one of the carbons in the scissile β-1,4-glycosidic bonds, i.e., oxidation of C1 and/or C4 of the glucose units, leading to carboxylic acid and/or keto functions at the cellulose surface [9, 11, 12]. LPMOs are widespread in the fungal kingdom, with five families of LPMOs (AA9, AA11, AA13, AA14, AA16) described in the CAZy database (www.cazy.org) [13, 14]. All charac-terized LPMOs that belong to the AA9 family are able to oxidatively cleave cellulose [15–18], and recent stud-ies have focused on their use to defibrillate cellulose and facilitate the production of nanocelluloses [19–21].

The ascomycete Podospora anserina has been studied for its impressive array of CAZymes involved in both cel-lulose and hemicelluloses breakdown, making it a model of choice to better understand the enzymatic decon-struction of plant biomass [22, 23]. Its genome encodes 33 AA9 LPMOs (PaLPMO9), eight of which contain a family 1 carbohydrate binding module (CBM1)-targeting cellulose. In the secretomes of P. anserina after growth on biomass, seven AA9 LPMOs were identified, five of which present a CBM1 [24]. Biochemical characteriza-tion of these enzymes showed various degrees of activity on cellulose, with higher total release of oxidized oligo-saccharides from cellulose for PaLPMO9A, PaLPMO9E and PaLPMO9H, all of which harbor a CBM1 module [17, 18]. PaLPMO9H was then further investigated for its capacity to disrupt cellulose fibers [19] and was shown to cleave mixed-linkage glucans, xyloglucan and glucoman-nan [25], and oligosaccharides [18]. Mass spectrometry analysis of the released products revealed that PaLP-MO9H catalyzes C4 oxidative cleavage of mixed-linkage

glucans and mixed C1/C4 oxidative cleavage of cellulose, glucomannan and xyloglucan [18, 25].

As stated earlier for P. anserina, the expansion in genes encoding AA9s has been observed in many fungal genomes. This gene multiplicity raises the question of the functional relevance at the organism level, i.e., functional redundancy or functional diversification and/or adapta-tions to substrate. Modular AA9 LPMOs bearing a CBM1 at their C-terminus are often predominantly secreted by filamentous fungi under lignocellulolytic conditions [26], but the role of these modules attached to LPMOs is not clearly established.

The roles of CBMs in glycoside hydrolase function have been widely explored (see [27] for review). Indeed, many glycoside hydrolases that attack the plant cell wall contain non-catalytic CBMs, which were first identified in cellulases [28]. CBMs are grouped into three types: type-A CBMs bind crystalline ligands while types B and C bind internal or terminal regions of polysaccharides, respectively. CBM1 is a type-A CBM, which binds crys-talline substrates using a planar surface [29]. CBMs not only target the enzymes to their substrates to promote catalysis [30, 31], but sometimes they can also modulate enzyme specificity [32]. CBMs are devoid of catalytic activity, but some studies suggest they play a role in the amorphization of cellulose through non-hydrolytic dis-ruption of the crystalline structure of cellulose [33, 34]. CBM1 appended to AA9 LPMOs may influence substrate binding, enzyme activity and/or regioselectivity, but the data are scarce and reported observations are contradic-tory. For instance, deletion of the CBM1 of NcLPMO9C had no effect on the degradation of PASC [35], whereas removal of the natural CBM from cellulose-active bacte-rial LPMOs abolished their activity [36].

Here, we investigate the role played by the CBM1 mod-ule to the cellulolytic activity of a fungal AA9 LPMO. PaLPMO9H was chosen as our model enzyme. The CBM1 module was truncated, and enzymatic activity was investigated using complementary approaches to exam-ine the release of soluble products and the cellulosic fib-ers themselves.

ResultsProduction of PaLPMO9H with and without CBM1To gain insight into the contribution of the CBM1 to the catalytic function of AA9 LPMOs, we selected PaLP-MO9H based on the previous biochemical analyses [18, 19, 25]. PaLPMO9H is a modular enzyme with two domains containing an N-terminal catalytic AA9 domain (16–243) and a C-terminal CBM1 domain (271–307) (Fig.  1). These two domains are connected through a serine/threonine/asparagine-rich linker comprising 27 amino acid residues. When the PaLPMO9H enzyme was

Page 3 of 10Chalak et al. Biotechnol Biofuels (2019) 12:206

truncated right after the catalytic module at position 244, we were unable to successfully produce the correspond-ing recombinant protein in P. pastoris (data not shown). Therefore, we decided to leave 16 amino acid residues of the linker to promote production of the recombi-nant enzyme. Using this strategy, we successfully pro-duced the CBM1-free PaLPMO9H enzyme truncated at position 259. In the rest of the study, the PaLPMO9H with the CBM1 is named LPMO-FL (full-length), and the PaLPMO9H without the CBM1 is named LPMO-CD (catalytic domain). As expected, deletion of the CBM1 decreased the enzyme’s molecular mass from ~ 38  kDa (LPMO-FL) to ~ 33  kDa (LPMO-CD). Appar-ent molecular mass was still slightly higher than theoreti-cal molecular mass (25.7  kDa) due to predicted O- and N-glycosylations (Additional file  1: Figure S1). LPMOs

are copper-dependent enzymes, which makes it crucial to check the correct copper protein loading. The amount of copper in each enzyme was quantified using induc-tively coupled plasma mass spectrometry (ICP-MS). Both enzymes were equally loaded with ~ 1 copper atom per molecule (i.e. 10.3 and 10.8  µM of Cu2+ for 10  µM of LPMO-FL and LPMO-CD, respectively).

Absence of CBM1 alters LPMO cellulolytic activity at low substrate concentrationThe action of LPMO-FL was first evaluated on three dif-ferent types of cellulose, i.e., phosphoric acid-swollen cellulose (PASC), nanofibrillated cellulose (NFC), and bacterial microcrystalline cellulose (BMCC) (Fig.  2a). As previously shown, LPMO-FL released both C4-oxi-dized (C4ox) and non-oxidized oligosaccharides from

Catalytic module

LPMO-FL

LPMO-CD

linker

Cellulose bindingmodule

AA9 CBM11 16 243 270 271 307

1 16 243 259AA9

(His)6

(His)6

Fig. 1 Schematic representation of the enzymes used in this study. LPMO‑FL (full‑length) and LPMO‑CD (catalytic domain) with amino‑acid numbering of the limits of each domain

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400

600

Glc

2

Glc

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lc5 C4ox C4ox C1-C4ox

Glc2-Glc6

Fig. 2 Analysis of soluble degradation products. a Products generated by LPMO‑FL upon degradation of 0.1% PASC, NFC or BMCC with 4.4 µM of LPMO in the presence of 1 mM of l‑cysteine, at 50 °C for 16 h. b Analysis of soluble degradation products generated by LPMO‑FL and LPMO‑CD upon degradation of 0.1% PASC with 4.4 µM of LPMO in the presence of 1 mM of l‑cysteine, at 50 °C for 4 h

Page 4 of 10Chalak et al. Biotechnol Biofuels (2019) 12:206

PASC [18]. However, using NFC as a substrate led to less products released, and using a more recalcitrant crystal-line substrate (BMCC) led to barely detectable products (Fig. 2a). We then compared the action of both LPMO-FL and LPMO-CD by measuring the release of sugars from PASC (Fig.  2b). LPMO-FL released higher amounts of soluble sugars (both oxidized and non-oxidized oligosac-charides) compared to LPMO-CD where soluble sugars were barely detectable (Fig. 2b).

Since LPMO-FL is active on soluble oligosaccharides [18], we investigated the activity of both LPMO-FL and LPMO-CD on cellohexaose as substrate (Additional file  1: Figure S2). A time-course analysis revealed that both enzymes were able to cleave cellohexaose, lead-ing mainly to Glc3 and Glc4 non-oxidized products and C4-oxidized products. Although LPMO-FL showed slightly better activity than LPMO-CD over the 24-h time-course, the observed cleavage of cellohexaose by LPMO-CD confirms that the enzyme lacking the CBM1 module is still functional.

LPMO-FL and LPMO-CD binding to PASC, BMCC and NFC was assessed in the absence of reductant using pull-down assays to assess the impact of the CBM1 (Additional file  1: Figure S3). LPMO-FL was observed in the bound fraction of all three cellulosic substrates tested. However, in the absence of CBM1, there were no bands corresponding to LPMO-CD in the bound frac-tion. Therefore, the CBM1 promotes LPMO binding to the cellulosic substrates.

Combined action of LPMO‑FL and LPMO‑CD with a cellobiohydrolaseTo assess the impact of LPMO-CD on cellulosic sub-strates, we assayed LPMO-FL and LPMO-CD enzymes in combination with the reducing end-acting cello-biohydrolase (family GH7 CBH-I) from T. reesei. Cellu-losic substrates were sequentially pretreated with either LPMO-FL or LPMO-CD before addition of the CBH-I enzyme. As both LPMOs and CBH-I act on soluble sub-strates, we implemented an LPMO post-treatment wash-ing step to assess the impact of LPMO treatment only on the insoluble fibers. LPMO pretreatment was beneficial on PASC and NFC but had no visible effect on the crys-talline substrate BMCC (Fig. 3). Pretreatment with either LPMO-CD or LPMO-FL increased cellobiose release from NFC substrate by approximately 30%. However, LPMO-FL pretreatment was more efficient on PASC sub-strate (60% increase in cellobiose production) compared to the LPMO-CD. Taken together, these results show that neither of the two LPMOs targets the crystalline fraction of cellulose. We believe that both LPMOs target amor-phous regions thus facilitating CBH-I activity on crys-talline cellulose. Moreover, under these experimental

conditions the presence of the CBM1 module is not strictly required for LPMO action.

Increasing substrate concentration reduces the need for the CBM1The next step was to assess whether substrate concen-tration has an influence on the activity of the enzymes. We increased the substrate concentration to 1% (w/v) to foster the probability of enzyme–substrate interactions in a CBM-free context. At high substrate concentration, soluble sugars released by LPMO-CD became detectable and were in the same range as soluble sugars released by LPMO-FL from PASC (Fig. 4). Interestingly, C1-oxidized (C1ox) products (Glc2ox-Glc4ox), which were barely detectable using LPMO-FL, were abundantly released by LPMO-CD (Fig.  4). The C4-oxidized products elut-ing at around 30  min were less abundant whereas the C1/C4-oxidized products eluting between 41 and 42 min were slightly increased. The absence of the CBM induced a modification of the regioselectivity pattern of the enzyme (Fig. 4).

Impact of LPMO on the insoluble fractionIn an effort to gain more insight into the role of CBM on the action of LPMOs, we evaluated the changes in mor-phology of kraft fibers in response to incubation with LPMO. First, we investigated fiber structure using opti-cal microscopy. Original kraft fibers are tens of microm-eters in diameter and around 1 mm long (Fig. 5a). After LPMO treatment, there were no visible changes in physi-cal appearance of the fibers, i.e., fibrous morphology or dimensions, in LPMO-FL (Fig.  5b) and LPMO-CD-treated samples (Fig.  5c). As previously described [19],

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the action of LPMOs alone does not produce a notice-able disintegration of kraft fibers, similarly to the action of cellulases [37–39]. Therefore, after LPMO treatment, fibers were mechanically dispersed then subjected to a short ultrasound treatment. Dispersion revealed the effect of LPMO on kraft fibers. Control samples showed some slight defibrillation whereas both LPMO-treated samples showed clear cell wall delamination (Fig. 5d–f). Both LPMO-FL and LPMO-CD seemed to create weak points within the fiber that facilitated the mechanical disintegration. To get a better picture of the effect of the LPMOs, we used atomic force microscopy (AFM) to ana-lyze samples (Fig. 5g–i). Topography images showed the presence of large fibers in control samples and a clear dis-sociation in LPMO-treated samples. LPMO-FL produced fibrillation of kraft fibers, forming an entangled network of ~ 5  nm-diameter nanofibrils. LPMO-CD also pro-duced a network of disintegrated fibers, but with thicker fiber bundles. Comparing the appearance of the fibers treated with LPMO-FL or LPMO-CD against controls, it is evident that both enzymes influence the cohesion and

architecture of the fibers, making them more prone to the mechanical forces caused by the dispersion. Based on AFM images, both LPMOs reduced fiber cohesion, but the presence of CBM appeared to enable LPMO-FL to defibrillate cellulose.

DiscussionThe functional relevance of CBMs and their contribution to the activity of the LPMO enzymes have already been investigated [36, 40], but in several cases, analysis surpris-ingly found modest and contradictory effects on enzyme activity. The role of CBMs appended to glycoside hydro-lases has been explored in depth [27], and it is generally acknowledged that the presence of CBMs increases the concentration of proteins at the surface of the substrate, thus increasing the activity of the enzyme [41]. Removal of the CBM attached to cellulases dramatically decreases the activity on insoluble substrates but not on soluble substrates [37, 42, 43]. A similar pattern was observed here with PaLPMO9H, as loss of the CBM dramati-cally affected the release of soluble sugars from cellulose

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Fig. 4 Analysis of degradation products generated by LPMO‑FL and LPMO‑CD. HPAEC chromatograms of the oligosaccharides released upon degradation of PASC [1% (w/v)] with 4.4 µM of LPMO in the presence of 1 mM of l‑cysteine, at 50 °C for 16 h. The sum of C1‑oxidized (C1ox) and C4‑oxidized (C4ox) oligosaccharides is indicated in the inset. *Glc2ox co‑eluted with Glc6

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whereas activity was retained on soluble cellooligosac-charides. However, when the substrate concentration of cellulose was increased, the lack of CBM did not seem to impede the action of PaLPMO9H (LPMO-CD), and solu-ble products were detected in the same range as the full-length enzyme. A similar pattern of action was observed with cellulases where a reduced amount of water coun-terbalanced the need for CBMs [44]. Our results are in line with the hypotheses drawn by Courtade et  al. [45] on a cellulose-active AA10 LPMO. Indeed, multiple cleavages are needed at the cellulose surface to release enough soluble products that can then be detected by ion chromatography. Here, we observed that the CBM1 appended to an AA9 LPMO promotes binding to cellu-lose and anchors the enzyme to the substrate, facilitating multiple localized cleavages. Conversely, AA9 LPMOs lacking CBM1 only weakly bind to cellulose and may thus

have a more random action on cellulose, thus limiting the number of localized cleavages and therefore the release of soluble cellooligosaccharides (<Glc6). This hypothesis is further supported by the synergistic effect with a CBH observed for both enzymes (with and without CBMs) on cellulose fibers and their capacity to defibrillate cellu-lose. Note, however, that although LPMO-FL was able to bind crystalline cellulose, it showed no detectable activ-ity, meaning that PaLPMO9H may target less-organized regions of cellulose, as already hypothesized in [19].

Surprisingly, CBM deletion was found to modify the regioselectivity pattern of the enzyme. The regioselec-tivity pattern was also modified when aromatic residues at the substrate-binding interface of HjLPMO9A were mutated [40], but removal of the HjLPMO9A CBM did not alter the regioselectivity of the enzyme even though the effect of mutations was increased in a CBM-free

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Fig. 5 Morphology of LPMO‑treated kraft fibers. Optical microscopy images of kraft fibers before (a–c) and after (d–f) mechanical dispersion for control samples (a, d), LPMO‑FL‑treated fibers (b, e) and LPMO‑CD‑treated fibers (c, f). AFM topographical images after LPMO treatment and dispersion for control kraft fibers (g), LPMO‑FL‑treated fibers (h) and LPMO‑CD‑treated fibers (i)

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context [40]. It seems that altering the mode of LPMO-to-substrate binding may slightly modify the position of the enzyme at the cellulose surface, thus generating a mixture of C1 and C4 cleavages. The fact that presence of the CBM may influence the regioselectivity of cellulose cleavage in LPMOs challenges the established pseudo-classification [46] that contains many exceptions, and raises questions as to the functional relevance of the C1 and/or C4 cleavage in LPMOs.

ConclusionsAssays of LPMO activity based on detecting soluble products warrant careful assessment taking into account the nature and concentration of the substrate. More generally, from a microbial degradation perspective, the fact that filamentous fungi secreted a wide range of AA9 LPMOs with and without CBMs may be exploitable to promote degradation depending on the substrate consist-ency. From a biotech perspective, the presence of a CBM appended to LPMOs could be mobilized to select targets for cellulose degradation purposes. However, regard-ing cellulose defibrillation for nanocellulose production, more work is needed to pinpoint the influence of the CBM on the efficiency of LPMOs used in the process.

Materials and methodsSubstratesThis study used several cellulosic substrates, represent-ing either the crystalline, amorphous, or alternating crystalline and amorphous regions, or quasi-natural fib-ers like the kraft fibers. Phosphoric acid-swollen cellu-lose (PASC) was prepared as described previously [18]. Nanofibrillated cellulose (NFC) obtained via endoglu-canase pretreatment followed by microfluidization was kindly provided by the Centre Technique du Papier (CTP, Grenoble, France). Bacterial microcrystalline cellulose (BMCC) was obtained from nata de coco cubes that were subjected to hydrochloric acid (2.5 N) hydrolysis at a temperature of 72 °C in three consecutive steps over a total time of 2  h, then separated by filtration and three centrifugation cycles at 10,000g for 10 min in which the acid supernatant was repeatedly replaced by water. Then, dialysis was done against distilled water. Bleached soft-wood kraft pulp was used as a substrate. Cellulose fibers were dispersed in 50  mM of sodium acetate buffer (pH 5.2) and stirred for 48 h prior to enzymatic assays [19].

Recombinant production of LPMO enzymesPaLPMO9H (protein ID CAP 61476) was produced in Pichia pastoris as described in [18]. To produce PaLP-MO9H without CBM, the gene region coding for its amino acid sequence 1–259 (see Fig.  1) was amplified and inserted into the pPICZalphaA vector (Invitrogen,

Cergy-Pontoise, France) using BstBI and XbaI restric-tion sites in-frame with the (His)6 tag. P. pastoris strain X33 and the pPICZalphaA vector are components of the P. pastoris Easy select expression system (Invitro-gen). All media and protocols are described in the Pichia expression manual (Invitrogen). Recombinant expression plasmids were sequenced to check the integrity of the corresponding sequences.

Transformation of competent P pastoris X33 was performed by electroporation with PmeI-linearized pPICZalphaA recombinant plasmid as described in [30]. Zeocin-resistant P. pastoris transformants were then screened for protein production. The best-pro-ducing transformant was grown in 1  L of BMGY con-taining 1  mL  L−1 of Pichia trace minerals 4 (PTM4) salts (2  g  L−1 CuSO4·5H2O, 3  g  L−1 MnSO4·H2O, 0.2  g  L−1 Na2MoO4·2H2O, 0.02  g  L−1 H3BO3, 0.5  g  L−1 CaSO4·2H2O, 0.5  g  L−1 CaCl2, 12.5  g  L−1 ZnSO4·7H2O, 22 g L−1 FeSO4·7H2O, biotin 0.2 g L−1, H2SO4 1 mL L−1) in flasks shaken at 30 °C in an orbital shaker (200 rpm) for 16 h to reach an OD600 of 2–6. Expression was induced by transferring cells into 200  mL of BMMY contain-ing 1 mL L−1 of PTM4 salts at 20 °C in an orbital shaker (200 rpm) for another 3 days. Each day, the medium was supplemented with 3% (v/v) methanol.

Enzyme purificationAfter harvesting cells by centrifugation (2700g for 5 min, 4 °C), the supernatant was adjusted to pH 7.8 just before purification, filtered on 0.22-µm filters (Millipore, Molsheim, France), and loaded onto a 5-mL HiTrap HP column (GE Healthcare, Buc, France) equilibrated with buffer A (Tris–HCl 50 mM pH 7.8, NaCl 150 mM, imi-dazole 10  mM) that was connected to an Äkta purifier 100 system (GE Healthcare). Each (His)6-tagged recom-binant enzyme was eluted with buffer B (Tris–HCl 50 mM pH 7.8, NaCl 150 mM, imidazole 500 mM). Frac-tions containing recombinant enzymes were pooled and concentrated with a 10-kDa vivaspin ultrafiltration unit (Sartorius, Palaiseau, France) and filter dialyzed against sodium acetate buffer 50 mM, pH 5.2. The concentrated proteins were incubated overnight with an equimolar equivalent of CuSO4 in a cold room and buffer exchanged in 50 mM sodium acetate buffer pH 5.2 using extensive washing in a 10-kDa ultrafiltration unit to remove traces of CuSO4.

Protein analysisProteins were loaded onto 10% Tris–glycine precast SDS-PAGE gels (BioRad, Marnes-la Coquette, France) and stained with Coomassie Blue. The molecular mass under denaturing conditions was determined with PageRuler Prestained Protein Ladder (Thermo Fisher Scientific,

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IL). The protein concentrations were determined by adsorption at 280 nm using a Nanodrop ND-2000 spec-trophotometer (Thermo Fisher Scientific) with theoreti-cal molecular masses and molar extinction coefficient derived from the sequences (49,640 and 39,545 M−1 cm−1 for LPMO-FL and LPMO-CD, respectively, measured at 280 nm in water).

ICP‑MS analysisThe ICP-MS analysis was performed as described in [47]. The samples were mineralized, then diluted in ultrapure water, and analyzed on an ICAP Q apparatus (Thermo Electron, Les Ulis, France). Copper concentration was determined using Plasmalab (Thermo Electron) software, at m/z = 63.

Qualitative cellulose‑binding assaysThe reaction mixtures were carried out at 0.3% (w/v) insoluble substrate loading (BMCC; NFC; PASC) and 30 µg of proteins were added. The reactions were done in 50 mM sodium acetate buffer pH 5.2 in a final volume of 200 µL without any l-cysteine addition. The tubes were incubated on ice for 1 h with gentle mixing every 10 min. After centrifugation at 14,000g for 10 min, the superna-tant (containing the unbound proteins) was carefully removed, then the polysaccharide pellets were washed twice (wash 1 and wash 2) by resuspending in buffer and centrifuged at 14,000g for 10 min. This step was repeated twice. The remaining pellet was finally resuspended in SDS-loading buffer without dye (with a volume equiva-lent to the unbound fraction removed) and boiled for 10 min to dissociate any bound protein. Unbound, wash 2 and bound fractions (45  µL supplemented with 5  µL of β-mercaptoethanol) were analyzed by SDS-PAGE to detect the presence or absence of the protein. The super-natant was recovered (supernatant 2: bound fraction), and 45  µL of supernatant 1 (unbound fraction), wash 2 and supernatant 2 (bound fraction) were analyzed by SDS-PAGE to detect the presence or absence of the pro-tein. We ran a control sample without any substrate to compare the results.

Enzymatic treatment of the substrates for the analysis of soluble sugarsAll the cleavage assays (on a final volume of 300 μL) con-tained 0.1% (w/v) of substrate (PASC, BMCC, NFC), 4.4 µM of PaLPMO9s, and 1 mM of l-cysteine, in 50 mM sodium acetate buffer pH 5.2. The enzymatic reactions were incubated in a thermomixer (Eppendorf, Montes-son, France) at 50 °C and 850 rpm for 16 h. At the end of the reaction, the samples were boiled at 100 °C for 15 min and then centrifuged at 15,000g for 10  min to separate the soluble and insoluble fractions. Assays at 1% (w/v)

PASC concentration were also done in the conditions mentioned earlier.

Combined assaysThe LPMO enzymatic assays were carried out sequen-tially with a cellobiohydrolase from T. reesei (CBH-I) as described in [48]. Assays were performed in a total vol-ume of 800 µL containing 0.1% (v/w) cellulose in 50 mM pH 5.2 acetate buffer with 8  µg of LPMO enzyme and 1  mM  l-cysteine. The samples were incubated in tripli-cate in a thermomixer (Eppendorf ) at 45 °C and 850 rpm, for 24 h. The samples were then boiled for at least 10 min and centrifuged at 15,000g for 10  min. The supernatant was removed, and the remaining insoluble fraction of the substrate was washed twice in buffer. Hydrolysis by CBH-I (0.8 µg) was performed in 800 µL of 50 mM pH 5.2 acetate buffer for 2 h at 45 °C and 850 rpm. The solu-ble fraction was analyzed as described below.

Analysis of oligosaccharidesOxidized and non-oxidized cellooligosaccharides gener-ated after LPMO action were analyzed by high-perfor-mance anion-exchange chromatography coupled with pulsed amperometric detection (HPAEC-PAD) (Ther-moFischer Scientific, IL) using a CarboPac™ PA1 col-umn (2 × 250  mm) and CarboPac™ PA1 guard column (2 × 50 mm) at a 0.25 mL min−1 flow rate as in [49]. Non-oxidized oligosaccharides were used as standards (Mega-zyme, Wicklow, Ireland).

Enzymatic treatment of the softwood pulp for the analysis of the insoluble fibersKraft fibers (100  mg) were adjusted to pH 5.2 with sodium acetate buffer (50  mM) in a final reaction vol-ume of 20  mL with 1  mM l-cysteine. Purified LPMO enzyme was added to the substrate at a final concentra-tion of 1.6 µM. Enzymatic incubation was performed at 50  °C under mild agitation for 16 h. Samples were then dispersed with a Polytron PT 2100 homogenizer (Kine-matica AG, Germany) for 3 min then ultrasonicated with a QSonica Q700 sonicator (20 kHz, QSonica LLC, New-town, CT) at 350 W ultrasound power for 3 min. The ref-erence sample was submitted to the same treatment but did not contain the LPMO enzyme.

Optical microscopyKraft fibers (reference and LPMO-treated) were depos-ited onto a glass slide and observed under a BX51 polar-izing microscope (Olympus France S.A.S.) with a 4× objective. Images (N ≥ 20) were captured by a U-CMAD3 camera (Olympus, Japan). The concentration of the fibers used was 2.5 g L−1 to visualize individual and separated fibers.

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Atomic force microscopy (AFM)Fiber dispersions were diluted at 0.1 g L−1. Samples were dialyzed against ultrapure water (spectral por-; molecu-lar porous membrane tubing 12–14  kDa) for 3  days to remove buffer, cysteine and released soluble sugars. They were later deposited onto mica substrates, allowed to set-tle for 45 min, and dried with Whatman filter paper. The final drying step was done in an incubator at 40  °C for 10 min before transfer to the AFM system. Topographical images on mica were registered by an Innova AFM sys-tem (Bruker). The images were collected in tapping mode under ambient air conditions (temperature and relative humidity) using a monolithic silicon tip (FESPA-V2) with a spring constant of 2.8 N m−1 and a nominal frequency of 75 kHz. Image processing was performed using WSxM 4.0 software. A series of reference images (between 3 and 11) were recorded to ensure the homogeneity of the sample.

Supplementary informationSupplementary information accompanies this paper at https ://doi.org/10.1186/s1306 8‑019‑1548‑y.

Additional file 1: Figure S1. SDS‑PAGE analysis of LPMO‑FL and LPMO‑CD purified enzymes. Ladder size is indicated in kDa. Figure S2. Time‑course analysis and quantification of the Glc4 (plain lines) and Glc3 (dotted lines) released by LPMO‑FL (triangles) and LPMO‑CD (circles) acting on cellohexaose over a total period of 24 h. Figure S3. Qualitative cellulose binding assays. Binding of LPMO‑FL and LPMO‑CD to (a) PASC (0.3% (w/v)), (b) NFC (0.3% (w/v)) and (c) BMCC (0.3% (w/v)). Lane 1, con‑trol (no substrate); lane 2, unbound material; lane 3, wash 1; lane 4, bound fraction. Experiments were carried out on ice using 30 µg of proteins and 50 mM sodium acetate buffer pH 5.2 in a final volume 200 µL without added l‑cysteine. Ladder size is given in kDa.

AbbreviationsAA: auxiliary activity enzyme; BMCC: bacterial microcrystalline cellulose; CAZyme: carbohydrate‑active enzyme; CBH: cellobiohydrolase; CBM: carbohydrate‑binding module; C1ox: C1‑oxidized oligos; C4ox: C4‑oxidized oligos; Glc2: cellobiose; Glc3: cellotriose; Glc4: cellotetraose; Glc5: cellopen‑taose; Glc6: cellohexaose; HPAEC‑PAD: high‑performance anion‑exchange chromatography coupled with amperometric detection; ICP‑MS: inductively coupled plasma mass spectrometry; LPMO: lytic polysaccharide monooxy‑genase; LPMO‑FL: LPMO full‑length; LPMO‑CD: LPMO catalytic domain; NFC: nanofibrillated cellulose; PASC: phosphoric acid‑swollen cellulose; SDS‑PAGE: sodium dodecyl sulfate‑polyacrylamide gel electrophoresis.

AcknowledgementsThe authors thank Florence Chaspoul from IMBE (CNRS Marseille) for ICP‑MS analysis.

Authors’ contributionsAC, MH and SG produced recombinant enzymes. AC and SG performed sub‑strate degradation tests and HPAEC–PAD analyses. AC, IHG and AL performed combined enzyme assays. AC, AV, CM and AD performed microscopy studies. JGB, AV and BC supervised work. AC and JGB wrote the manuscript with input from AV and BC. All authors reviewed the manuscript. Figures were prepared by AC, SG and AV. All authors read and approved the final manuscript.

FundingThis study was funded by the “Region Pays de la Loire” and an INRA Ph.D. grant to AC.

Availability of data and materialsThe datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Ethics approval and consent to participateNot applicable.

Consent for publicationNot applicable.

Competing interestsThe authors declare that they have no competing interests.

Author details1 Biopolymères Interactions Assemblages, INRA, Nantes, France. 2 Biodiver‑sité et Biotechnologie Fongiques, UMR1163, INRA, Aix Marseille Université, Marseille, France.

Received: 10 May 2019 Accepted: 24 August 2019

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Titre : Évaluation de l'activité d'une LPMO AA9 modulaire sur des substrats cellulosiques.

Mots clés : Podospora anserina, cellulose, LPMO9H, « carbohydrate binding module », CBM1.

Contexte: Les « lytic polysaccharides monooxygenases »(LPMO), sécrétées par les champignons filamenteux, jouent un rôle important dans la dégradation de la biomasse lignocellulosique récalcitrante. Ils peuvent se présenter sous la forme de protéines multidomaines fusionnées à un « Carbohydrate Binding Module » (CBM). Sur le plan biotechnologique, les LPMOs sont des outils prometteurs et innovants pour la production de nanocelluloses et de biocarburants, mais leur action directe sur les substrats cellulosiques n’est pas pleinement comprise. Résultats: Nous avons étudié l'action d’un module de liaison à la cellulose (CBM1) lié à la LPMO9H de Podospora anserina (PaLPMO9H) en utilisant des substrats cellulosiques modèles. La suppression du CBM1 a affaibli la liaison à la cellulose nanofibrillée, à la cellulose amorphe et cristalline. Bien que la libération de sucres solubles ait été considérablement réduite dans des conditions normales, la LPMO tronquée a conservé une certaine activité sur les

oligosaccharides solubles. L'action cellulolytique de la LPMO tronquée a été démontrée à l'aide d'expériences de synergie avec une cellobiohydrolase (CBH). En effet, la LPMO tronquée était encore capable d’améliorer l’action du CBH sur la cellulose. L'analyse de la fraction insoluble a confirmé que le module CBM1 n'était pas strictement nécessaire pour favoriser la perturbation du réseau de cellulose. Sur la base de ces résultats, nous avons réduit la quantité d'eau dans la réaction pour augmenter la probabilité d'interaction enzyme-substrat dans un contexte sans CBM. Ces conditions ont amélioré les performances de PaLPMO9H sans CBM en termes de libération de produits. Il est intéressant de noter que l’élimination du CBM a modifié la régiosélectivité de PaLPMO9H avec une libération importante de produits oxydés en C1. Ces résultats fournissent des informations sur le mécanisme d’action des LPMO fongiques sur la cellulose pour produire des nanocelluloses et des biocarburants.

Title Assessing the activity of a modular AA9 LPMO on cellulosic substrates

Keywords : Podospora anserina, cellulose LPMO9H, Carbohydrate Binding Module 1, CBM1

Background: Lytic polysaccharide monooxygenases (LPMOs) secreted by filamentous fungi play a key role in the degradation of recalcitrant lignocellulosic biomass. They can occur as multidomain proteins fused to a carbohydrate-binding module (CBM). On a biotechnological point of view, LPMOs are promising and innovative tools for the production of nanocelluloses and biofuels but their direct action on cellulosic substrates is not fully understood. Results: We probed the action of the family 1 CBM (CBM1) appended to the LPMO9H from Podospora anserina (PaLPMO9H) using model cellulosic substrates. As expected, the deletion of the CBM1 weakened the binding to nanofibrillated, amorphous and crystalline cellulose. Although the release of soluble sugars from cellulose was drastically reduced under standard conditions, the truncated LPMO retained activity on soluble oligosaccharides.

The cellulolytic action of the truncated LPMO was demonstrated by synergy experiments with a cellobiohydrolase (CBH). The truncated LPMO was still able to improve the efficiency of the CBH on cellulose nanofibrils similarly to the full length LPMO. Analysis of the insoluble fraction of cellulosic substrates confirmed that the CBM1 module was not strictly required to promote the disruption of the cellulose network. Based on these results we reduced the amount of water to increase the probability of enzyme-substrate interaction in a CBM-free context. These conditions enhanced the performance of PaLPMO9H without CBM in terms of products release. Interestingly, removing the CBM altered the regioselectivity of PaLPMO9H increasing C1 oxidized products. These results provide insights into the mechanism of action of fungal LPMOs on cellulose to produce nanocelluloses and biofuels.