Abstract
Polysaccharides from plant biomass are the most abundant renewable chemicals on Earth and can potentially be converted to a wide variety of useful glycoconjugates. Potential applications of glycoconjugates include therapeutics and drug delivery, vaccine development and as fine chemicals. While anomeric hydroxyl groups of carbohydrates are amenable to a variety of useful chemical modifications, selective cross-coupling to non-reducing ends has remained challenging. Several lytic polysaccharide monooxygenases (LPMOs), powerful enzymes known for their application in cellulose degradation, specifically oxidize non-reducing ends, introducing carbonyl groups that can be utilized for chemical coupling. This study provides a simple and highly specific approach to produce oxime-based glycoconjugates from LPMO-functionalized oligosaccharides. The products are evaluated by HPLC, mass spectrometry and NMR. Furthermore, we demonstrate potential biodegradability of these glycoconjugates using selective enzymes.
Similar content being viewed by others
Introduction
Over the years, numerous methods for synthesis of glycoconjugates have been developed, mainly using traditional synthetic methods. An unprotected carbohydrate contains multiple hydroxyl groups with similar reactivity. Coupling ligands to a specific position is therefore challenging and normally requires an exhaustive number of chemicals and synthetic steps1,2,3. This is one of the major hurdles for efficient utilization of some of Nature’s most abundant molecules as building blocks for novel biomolecules. Glycoconjugates have a considerable potential as drugs, fine chemicals and in material science4. Moreover, the efficiency of enzymatic depolymerization of plant polysaccharides has improved tremendously in recent years, partly due to the inclusion of oxidative enzymes in enzyme mixtures5,6,7. This development enables more efficient production of plant derived carbohydrate-based “bulk chemicals”, including a wide variety of structurally diverse oligosaccharides that could be converted to glycoconjugates.
Oxidative carbohydrate-active enzymes are involved in a multitude of biological processes, including biomass decay. These enzymes use different mechanisms to generate oxidations on mono-, oligo- and/or polysaccharides. These oxidation reactions produce either carbonyls or carboxylic acids and, for poly- and oligomeric substrates, these reactions involve enzymes such as LPMOs5,8,9,10, cellobiose dehydrogenase (CDH)11,12, galactose oxidase13, pyranose dehydrogenase14,15 and glucooligosaccharide oxidases16. Together, these enzymes enable a range of oxidative functionalizations17 and offer several possibilities for subsequent glycoconjugation18. Notably, there are very few enzymes that oxidize the non-anomeric hydroxyls in the sugar rings of oligo- and polysaccharides.
LPMOs may play a special, albeit so far scarcely explored, role in glyco-functionalization, since they are active on polymers and can functionalize polymeric surfaces. The initial step of the lytic action of LPMOs on cellulose entails breaking the energetically strong C–H bond19 at position C1 or C4, catalyzed by a triangularly shaped catalytic copper site termed the “histidine brace”9. Some LPMOs oxidize C1, thus generating a carboxylic acid that could also be generated in alternative ways, for example using CDH. Interestingly, other LPMOs have the ability to specifically oxidize C4, generating a ketone functionality that is hard to generate in other ways. Due to the discovery of an increasing number of C4-oxidizing LPMOs active on abundant polysaccharides such as cellulose20, glucomannan21, xylan22 and xyloglucan21, several target substrates for non-reducing end functionalization by LPMOs are now available.
A feasible way to synthesize heteropolymers is via polycondensation (analogous to the synthesis of well-known materials like polyesters and polyurethanes). This requires carbohydrates that are bi-functionalized. While functionalization and/or cross coupling to carbohydrate reducing ends is relatively straightforward, doing the same at the non-reducing end has proven more challenging, although elegant approaches to this using galactose oxidase23 and recently galactose oxidase/pyranose dehydrogenase and transaminidase have been shown24. The possibility to specifically functionalize C4 using LPMOs opens a possible route towards generating other bi-functional carbohydrate building blocks.
Here we describe a highly selective chemo-enzymatic coupling procedure for the C4 position of oligosaccharides derived from one of the most abundant polysaccharides in Nature, cellulose (Fig. 1 sketches the different synthetic steps). We further demonstrate that this allows the production of enzymatically cleavable bi-functional glycoconjugates.
Results and discussion
Demonstrating the regioselectivity of NcLPMO9A
Several LPMOs characterized so far have mixed activities leading to a combination of C1- and C4-oxidized products, whereas some strict C4-oxidizers act on oligomeric substrates and thus produce short products25. These properties would interfere with the coupling chemistry and complicate analysis in this proof of concept study. Hence, we searched for an LPMO that is not active on oligosaccharides and only releases C4-oxidized products with a degree of polymerization (DP) that would provide oligosaccharides of a suitable range (3–6). Based on previous studies, the C4-oxidizing Neurospora crassa enzyme, NcLPMO9A (NCU02240)26,27, which is not active on soluble cellooligosaccharides, was selected. More detailed analysis, necessary to validate this enzyme’s suitability for the present study, showed that it releases longer, oxidized oligosaccharides (predominantly DP 3–5) (Fig. 2). Furthermore, MS/MS analysis displayed in Fig. S1 showed fragmentation patterns typical of C4-oxidized cello-oligosaccharides25.
When NcLPMO9A was combined with a cellobiose dehydrogenase (MtCDH from Myrococcum thermophilum)28, the dominating ketone signal in the MALDI-ToF MS spectra shifted by 16 or 38, representing the formation of an aldonic acid (+ 16), which subsequently has a tendency to form a double sodium adduct (M − H + 2Na; + 38) (Fig. 2). Since CDH oxidizes oligosaccharides in the reducing end, this observation confirms that NcLPMO9A oxidizes in the non-reducing end only.
Since it is challenging to determine the oxidative regioselectivity of LPMOs by HPAEC and MALDI29,30, we designed a precise and simple way for probing C4-oxidation on oligosaccharides, inspired by previous work by Beeson et al.20 who looked at monosaccharides. Reduction of the ketone at C4 results in a mixture of galactose and glucose and hence reduction of a C-4 oxidized cello-oligosaccharide will result in the formation of a mixture of oligosaccharides with either a glucosyl or a galactosyl at the non-reducing end and a glucitol (GlcOH) in the reducing end. To assess this in detail, we first generated GalGlcn (n = 3 or 4) standards (Fig. 3) by using UDP-Gal (donor), cellotriose/tetraose (acceptor) and a galactosyltransferase (Fig. 3E).
These oligosaccharides have considerably shorter elution times in HPAEC than their corresponding cello-oligosaccharides (Fig. 4A). Oligosaccharides generated by NcLPMO9A and our in-house prepared oligomeric standards were reduced to completion using NaBD4 and the resulting oligomeric products were analyzed directly by HPAEC and MALDI ToF MS (Figs. 2, 4B,C). Reaction products generated by NcLPMO9A indeed yielded a mixture of (Glc)nGlcOH and Gal(Glc)nGlcOH (n = 2–4) oligosaccharides confirming C4 oxidation. These results further prove that NcLPMO9A only oxidizes C4.
Notably, this reduction approach, used here to prove C4 oxidation, also provides a relatively simple method for quantification of oxidized products, based on using NaBD4 for reduction. After reduction, oligosaccharides with oxidized C4 will possess two deuteriums whereas non-oxidized oligosaccharides will only contain one deuterium, thus yielding an easily detectable difference of m/z = 1 (Fig. 4C). Internal standards could thus be produced from non-oxidized oligosaccharides as described above, possibly using NaBH4 for reduction, generating a more pronounced difference of m/z = 2.
Figures 1 and 3 further show that NcLPMO9A primarily releases oligosaccharides of DP3-5, which is a suitable size for the purpose of this study. Several experiments were performed in order to evaluate the specificity of NcLPMO9A. Screening of other substrates using MALDI-ToF MS for product detection showed that, in contrast to several other C4 oxidizing LPMO9s, NcLPMO9A is not active on curdlan [β-(1,3) glucan], pustulan [β-(1,6) glucan], β-chitin, glucomannan, xylan, xyloglucan, mixed linkage glucan [β-(1,3),(1,4)] and is thus highly specific for cellulose. Furthermore, there was no activity on cello-oligosaccharides with a degree of polymerization (DP) of 2–6, which shows that this enzyme is not active on solubilized oligomeric products. Together the properties demonstrated above makes NcLPMO9A suitable for this proof of concept study on C4-functionalized cellooligosaccharides.
Generation of mono- and difunctional glycoconjugates
Oxidized products generated by NcLPMO9A (C4-oxidized) or by a combination of NcLPMO9A and CDH (C1 and C4-oxidized; Fig. 1) were subjected to targeted carbonyl functionalization using a hetero-bi-functional linker (aminooxy-linker), 2-(aminooxy)-1-ethanaminium dichloride, to generate oximes (Fig. 5). Product formation was confirmed by MALDI-ToF MS (Fig. 5) and porous graphitized carbon electrospray ionization mass spectrometry (PGC-ESI-MS) (Fig. 6) as well as by NMR (Fig. 7).
Due to the presence of a carbonyl function in the reducing end, oxime formation takes place at both ends of the C4-oxidized oligosaccharide. Chromatography/tandem mass spectrometry (LC–MS/MS) with PGC yielded masses corresponding to three double-oxime functionalized oligosaccharides (m/z 619.25, 781.28 and 943.35). Furthermore, the obtained fragmentation patterns could be matched with the oxime functionalized oligosaccharides (Fig. 6). Notably, when analyzing the oxime functionality on PGC we observed a considerably increased retention of the oligosaccharides (Fig. 6), compared to the corresponding C-4 oxidized oligosaccharides30. This may be due to charge effects which are well known to increase the retention of charged solutes on PGC. Replacing ammonium acetate with a stronger eluent like formic acid in the gradient was needed to enable elution. Since the oxime linkers will occur in an equilibrium between charged and neutral states, this may have given rise to the peak broadening observed. Upon viewing the associated fragmentation patterns there were no signs of isomer formation as the fragmentations were consistent throughout the elution times for each particular peak (Fig. S2).
In double oxidized oligosaccharides, containing a CDH-generated carboxyl functionality that will block oxime formation at the former “reducing end”, only the carbonyl function at C4 in the non- reducing end will react. Indeed, for C4-oxidized oligosaccharides that were protected in their reducing end by CDH-catalyzed oxidation to an aldonic acid, only one oxime was formed, in the non-reducing end. The efficiency of the reaction was assessed by mass spectrometry (Figs. 4, 5) and the position of the oxime was verified by NMR both for the reducing and the non-reducing end (Fig. 7, Fig. S3, Table S1). Obviously, these double oxidized oligosaccharides allow the formation of hetero-linked compounds by utilizing both carbonyl specific (at the non-reducing end) and carboxyl-specific coupling reactions in the down-stream (former reducing) end.
Structural elucidation by NMR provided direct proof of the structure of the putatively bi-functionalized products generated by coupling aminooxy-linkers to either one or both ends oligosaccharides. The individual monosaccharides were assigned by starting at the anomeric signal as well as from the primary alcohol group at C6 and then following 1H–1H connectivity using DQF-COSY, H2BC and [1H–13C] HSQC-[1H,1H] TOCSY (full assignment of shift values in Table S1). Most of the carbon chemical shifts were obtained from [1H–13C] HSQC (Fig. 7).
The [1H–13C] HMBC spectrum provided long range bond correlations that connect the monosaccharides to the aminooxy-linker. The carbon chemical shift of the C4 oxidized end was determined by combining the correlations from a [1H–13C] HSQC spectrum and a [1H–13C] HMBC spectrum (see Fig. 7B). Both for the reducing end (Glc1–H/C-2) and for the C4 oxidized non-reducing end (C4ox-3 and C4ox-5) HMBC correlations are observed from proton signals 4.61 ppm and 5.01/4.27 ppm to a carbon signal at 155.4 ppm and 159.2 ppm, respectively (see Fig. 7B). These carbon chemical shifts fit well to the expected carbon chemical shifts for an aminooxy group, which would form upon coupling. Altogether, the NMR results suggest that the structure of the bi-functional products correspond with the structures depicted in Fig. 6C.
Enzyme degradability
To explore the potential of further expanding the functionality of the modified cello-oligosaccharides we tested various cellulase treatments. The aminooxy-functionalized oligosaccharides can indeed be enzymatically degraded using a cellulase. This indicates that glycoconjugates produced in this study may be biodegradable which is an important aspect in the applicability of such biobased chemicals, whilst it also allows for the formation of glycoconjugates with an unmodified reducing end This effectively expands the application potential of the present technology and underpins the flexibility of these conjugates, by allowing for the potential of a multistep approach to labeling. Two different cellulases were used to demonstrate degradability of the functionalized oligosaccharides (Fig. 8). Both Cel6B from T. fusca and Cel5A from H. jecorina degraded the aminooxy-functionalized oligosaccharides.
Both enzymes degraded bi-functionalized DP5 and DP6 completely, and Cel5A also degrade most of bi-functionalized DP4. Since the mass difference of degradation products with the oxime linker in the reducing end is m/z + 2 compared to products where the oxime linker occurs in the non-reducing end, the mass spectra reveal preferred sites for cellulase cleavage. For example, Cel5A primarily converted the double-functionalized pentamer to a trimer with C4-oxime linker (583.2) and a dimer with a reducing end linker (423.1), whereas conversion to a dimer with C4-linker (421.1) and a trimer with a reducing end linker (585.2) was less frequent. For Cel6B the degradation pattern for the double-functionalized pentamer was slightly different; here the main products were a trimer with a reducing end linker (585.2) and a dimer with a C4-linker (421.1). This exemplifies a rational approach for generating defined functionalized oligosaccharides. Essentially, by playing with the initial reaction (one or two oxime functionalizations) and the use of cellulases, in principle any combination of a natural and a modified chain end can be generated. More uniform products may be obtained by first purifying the C4-oxidized cello-oligosaccharides, which could be carried out using Porous Graphitized Carbon columns or HILIC30.
By combining C4-specific LPMOs and CDHs, more complex glycoconjugates may be generated since one may employ chemistries pertaining to carbonyl groups, such as the method described above, and chemistries pertaining to carboxyl groups, for example using carbodiimide activation31. Notably, C1-specific LPMOs, or even LPMOs with a mixed C1/C4 activity could also be considered for this purpose. Importantly, the use of LPMOs in principle allows partial surface oxidation of polysaccharide fibers32,33,34, which may enable functionalization of fiber surfaces without affecting the tensile strength of the fibers. Such applications could be of commercial interest.
Further derivatization of the amino groups introducing fluorescein isothiocyanate (FITC) handles
The functional group introduced through aminooxy functionalization was an amino group, and the usefulness of this group for further functionalization of the glycoconjugate was demonstrated by labeling with fluorescein isothiocyanate (FITC) that is highly reactive towards amines. Indeed, after FITC labeling, oligosaccharides with oxime functionalization in both ends carried either one (due to incomplete labeling) or two FITC molecules (Figs. 9A, 10).
On the other hand, labeling oligosaccharides oxidized by CDH and thus carrying only one oxime functionalization carried only a single FITC specifically labeled at C4 in the non-reducing end (Figs. 1, 9B). The aminooxy functionalization shown in this study demonstrated only one of many possibilities for functionalization. Other functional groups such as azides and alkynes are also feasible and will extend the potential of the approach described here.
Conclusion and implications
Potential applications of the C4-specific functionalization demonstrated here, include regiospecific immobilization of C4-linked oligosaccharides to solid surfaces such as microarrays, conjugation to protein carriers for immunization purposes, and C4-specific conjugation to nano- or microparticles, or quantum dots35. C4-specific functionalization also provides a potential novel analytical tool for studying whether glycosidases act at the reducing end or non-reducing end of oligosaccharides, a feature that has proven challenging to access and may require multistep approaches to resolve36. Generating oligosaccharides that are either blocked in the non-reducing end or the reducing end could, in a straightforward way, aid in providing unequivocal evidence of which end is targeted by a given glycosidase. Furthermore, FITC labeling has been shown useful for studying the uptake of specific carbohydrates by microbes37. The labeling strategy presented here provides new tools for such studies and could advance these by enabling studies of the directionality of glycan uptake.
Chemo-enzymatic protocols enable synthesis of glycoconjugates with unprecedented precision and are easier than the multistep procedures that are needed when using conventional carbohydrate chemistry. Much progress has been made in recent years on regioselective modification of unprotected sugars38, for example through selective chemical oxidation39, allowing increasingly efficient chemical synthesis of oligosaccharide-based glycoconjugates. The current study provides a novel chemo-enzymatic route towards glycoconjugates from oligo- and even polysaccharides, providing an attractive alternative to synthesis starting from monosaccharides40. We envisage that technologies such as the one presented in this study will help pave the way towards environmentally friendly valorization of biomass and production of useful biochemicals.
Materials and methods
Materials
Cellobiose, cellotriose, cellotetraose, cellopentaose and cellohexaose were from Megazyme (Bray, Ireland). Sodium cacodylate, manganese (II) chloride tetrahydrate, uridine diphosphate galactose (UDP-Gal), fluorescein isothiocyanate (isomer I) and galactosyl transferase from bovine milk were from Sigma. AlexaFluor488 C5-aminooxyacetamide, and bis(triethylammonium) salt were from Invitrogen (Nærum, Denmark). 2-(aminooxy)-1-ethanaminium dichloride was from ABCR GmbH (Karlsruhe, Germany). 2,5-Dihydroxy-benzoic acid was from Bruker Daltonics (Bremen, Germany). The LPMO used in this study, from Neurospora crassa (NcLPMO9A), and cellobiose dehydrogenase from Myrococcum thermophilum (MtCDH) were produced and purified according to Petrovic et al. 201926 and Flitsch et al. 201928, respectively. The endocellulase Cel5A from Hypocrea jecorina was produced according to Saloheimo et al. 198841 and the exocelluase Cel6B from Thermobifida fusca was produced according to Vuong and Wilson 200942.
Methods
Enzyme reactions
-
(1)
NcLPMO9A (1 µM), ascorbic acid (4 mM), phosphoric acid swollen cellulose (PASC, 1% w/v), TrisHCl (10 mM, pH 8.0), 50 °C, 1,000 rpm, 12 h.
-
(2)
MtCDH (1 µM), NcLPMO9A (1 µM), ascorbic acid (1 mM), phosphoric acid swollen cellulose (PASC, 1% w/v), 50 °C, 1,000 rpm, 12 h. MtCDH was used to form double oxidized products.
-
(3)
Cel5A (1 µM, endocellulase from Hypocrea jecorina), oxime functionalized oligosaccharides (1 mg/mL), 50 °C, 1,000 rpm, 12 h.
-
(4)
Cel6B (1 µM, exocellulase from Thermobifida fusca), oxime functionalized oligosaccharides (1 mg/mL) 50 °C, 1,000 rpm, 12 h.
Enzyme reactions (3) and (4) were used to demonstrate that oxime functionalized oligosaccharides were enzymatically degradable.
Production of galactosyl-cello-oligosaccharide standards
Galactosyl-cello-oligosaccharides were synthesized using a modified procedure from literature43. Briefly, 1 mg of cellotriose or cellotetraose was dissolved in 500 µL of 40 mM sodium cacodylate buffer pH 6.8 containing 40 mM MnCl2, 5 mg of UDP-Gal and 0.5 mg of galactosyl transferase from bovine milk (Sigma-Aldrich). The galactosyl transferase will add one galactose to the non-reducing end of the cello-oligosaccharides, forming a β-(1–4) glycosidic bond44. The reaction was left to proceed for 3 days at 37 °C with shaking at 1,400 rpm. The reaction mixture was left to cool down to room temperature and was directly applied to a CarboGraph column (Grace, Columbia, USA) for desalting of the oligosaccharides, essentially according to a previously published protocol45.
Functionalization of C4-oxidized cello-oligosaccharides with a hetero-bi-functional linker and labeling with a fluorophore, forming oxime linked products
5 mL of the supernatant from the reactions with PASC (10 mg/mL) prepared as described in enzyme reaction 1 above were used in further experiments. The oligosaccharides were functionalized by reacting carbonyl groups with 2-(aminooxy)-1-ethanaminium dichloride to form oximes, using a modified procedure from literature46. The reaction was conducted in 0.1 M NaOAc buffer, pH 4.9, with ca. 50 molar equivalents of the aminooxy linker based on the amount of oxidized oligosaccharides, over a course of 6 days at 37 °C with shaking at 1,400 rpm. The products were then purified using active carbon, essentially according to literature45. Oxime-functionalized oligosaccharides were labeled with fluorescein isothiocyanate (FITC, ca. 6 molar equivalents with respect to the amino-group content of the oligosaccharide) by incubation in 10 mM sodium bicarbonate buffer, pH 9.5, for 1 h, with shaking at 1,400 rpm, at room temperature and in the dark. The reaction mixture was then freeze-dried.
Reduction of soluble products generated by NcLPMO9A
Reduction of oligosaccharides was conducted by mixing 10 µL of oligosaccharide sample from reaction 1 with 65 µL H2O (MilliQ) containing 700 µg NaBH4 or NaBD4. The reduction reaction was incubated over night at ambient temperature and then quenched by adding 20 µL 25 mM sodium acetate. Chromatographic analysis by HPAEC (see below) was done with the sample as is, whereas a further sample preparation for MALDI-ToF-MS was done as follows. Approximately 10 µL of a 1:1 (w/w) suspension of H2O:Supelclean ENVI Carb (Sigma-Aldrich) was packed in a pipette tip containing a C8-disc that was trapping the Supelclean material. The bed was conditioned with 50 µL H2O. The sample (5 µL) was applied, rinsed with 50 µL H2O and then eluted with 20 µL acetonitrile. The eluate was then analyzed by MALDI ToF-MS.
HPLC analysis of oligosaccharides
Oligosaccharides were analyzed using three different chromatographical principles. The first principle, high-performance anion-exchange chromatography (HPAEC) at high pH and coupled with pulsed amperometric detection (PAD), was conducted on an ICS3000 system from Dionex (Sunnyvale, CA, USA), using previously described conditions and gradients30,47. The system was set up with PAD using disposable electrochemical gold electrodes. Two µL samples were injected on a CarboPac PA1 2 × 250 mm analytical column equipped with a CarboPac PA1 2 × 50 mm guard column and columns were kept at 30 °C.
The second principle, porous graphitic carbon (PGC) chromatography, was applied using an Ultimate 3000RS (Dionex) UHPLC system, which was connected in parallel (10:1 split) to an ESA Corona Ultra charged aerosol detector (ESA Inc.,Dionex, Sunnyvale, USA) and a Velos pro LTQ linear ion trap (Thermo Scientific, San Jose, CA USA). The PGC column (Hypercarb, 3 µm, 2.1 × 150 mm) including a guard column (Hypercarb, 3 µm, 2.1 × 10 mm) was operated at 70 °C using previously described conditions and gradients30. In some cases, 10 µM formic acid was used instead of ammonium acetate, to improve resolution of the oxime coupled products. Two µL samples were injected.
The third principle, hydrophilic interaction chromatography (HILIC), was applied using an Agilent 1290 Infinity (Agilent Technologies, Santa Clara, CA, USA) UHPLC system, which was connected in parallel (1:10 split) to an Agilent 1260 fluorescence detector—Ex 490 nm, Em 520 nm (Agilent Technologies, Santa Clara, CA, USA) and a Velos pro LTQ linear ion trap (Thermo Scientific, San Jose, CA, USA). The HILIC column (bioZen Glycan, 2.6 µm, 2.1 × 100 mm) including a guard column (SecurityGuard ULTRA with bioZen Glycan cartridge, 2.1 × 2 mm) was operated at 50 °C, running at 0.3 mL/min, and using 50 mM ammonium formate pH 4.4 (eluent A) and 100% acetonitrile (eluent B). Samples were eluted using the following gradient: initial starting ratio of 85% B and 15% A, gradient to 75% B and 25% A from 0 to 5 min, gradient to 60% B and 40% A from 5 to 25 min, gradient to 40% B and 60% A from 25 to 28 min, isocratic from 28 to 32 min, gradient to 85% B and 15% A from 32 to 35 min. Two µL of the samples were injected.
MALDI-ToF analysis of oligosaccharides
Two µL of a 9 mg/mL solution of 2,5-dihydroxybenzoic acid in 30% (v/v) acetonitrile was applied to an MTP 384 target plate ground steel TF (Bruker Daltonics). One µL of the sample was then mixed into the DHB droplet followed by drying under a stream of hot air. The samples were analyzed with an Ultraflex MALDI-ToF/ToF instrument (Bruker Daltonics GmbH, Bremen, Germany) with a Nitrogen 337 nm laser beam. The instrument was operated in positive acquisition mode and controlled by the FlexControl 3.3 software package. The acquisition range used was from m/z 200 to 7,000. The data were collected from averaging 250 laser shots, with the lowest laser energy necessary to obtain sufficient signal to noise ratios. Peak lists were generated from the MS spectra using Bruker FlexAnalysis software (Version 3.4).
Mass spectrometry of oligosaccharides, direct injection MS, PGC-MS and HILIC-FLD-MS
For direct injection, oligosaccharides were analyzed using an LTQ-Velos Pro linear ion trap mass spectrometer (Thermo Scientific, San Jose, CA, USA) connected to an Ultimate 3000RS HPLC (Dionex, Sunnyvale, CA, USA). The setup was used for direct injection without a column; the pump delivered 200 µL/min of 0.03 µM formic acid in 70% acetonitrile and data was acquired for 24 s after injection. For the MS, the capillary voltage was set to 3.5 kV and the scan range was m/z 150–2,000 using two micro scans. The automatic gain control was set to 10,000 charges and the maximum injection time was 20 ms. For fragmentation of selected precursor ions by MS/MS, the normalized collision energy was set to 37 and three micro scans were used. For PGC-MS, the same MS-parameters were used as for direct injection with the exception that the scan range was m/z 250–2,000. For HILIC-FLD-MS the instrument was operated in positive mode with an ionization voltage of 3.5 kV, auxiliary and sheath gas settings of 5 and 30 respectively (arbitrary units) and with capillary and source temperatures of 300 °C and 250 °C, respectively. The scan range was set to m/z 110–2,000 and MS/MS analysis was performed with CID fragmentation with helium as the collision gas. All data were recorded with Xcalibur version 2.2 (Thermo Scientific).
NMR sample preparation
Samples for NMR analysis were lyophilized, then dissolved in 1,000 µL 99.9% D2O (Chiron, Trondheim, Norway), frozen and lyophilized. The dried samples were then dissolved in 300 µL 99.9% D2O and transferred into a 4 mm Shigemi tube (Shigemi, Allison Park, PA, USA).
NMR data acquisition and analysis
All homo- and heteronuclear NMR experiments were carried out on a BRUKER Avance III HD 800 MHz spectrometer (Bruker BioSpin AG, Fällanden, Switzerland) equipped with a 5 mm TXI z-gradient probe.
For chemical shift assignment, the following spectra were recorded: 1D 1H, 2D double quantum filter correlated spectroscopy (DQF-COSY), 2D total correlation spectroscopy (TOCSY) with 70 ms of mixing time, 2D48 heteronuclear single quantum coherence (HSQC) with multiplicity editing, 2D48 heteronuclear 2-bond correlation (H2BC), 2D [1H–13C] HSQC-[1H,1H] TOCSY with 70 ms of mixing time on protons, and 2D [1H–13C] heteronuclear multiple-bond correlation (HMBC) with BIRD (bilinear rotation decoupling) filtering to suppress first order correlations. These spectra were all recorded at 25 °C. Diffusion-ordered spectroscopy (DOSY) was used to measure the diffusion of the coupled products. A 2D DOSY was set up using a Bruker BioSpin stimulated echo pulse sequence with bipolar gradients (STEBPGP). Gradient pulses of 2 ms duration (δ) and 32 different strengths varying linearly from 0.03 to 0.57 T⋅m−1 were applied and the diffusion delay (Δ) was set to 80 ms. The DOSY spectrum was recorded at 25 °C. The spectra were recorded, processed and analyzed using TopSpin 3.2 software (Bruker BioSpin AG).
References
Wang, C. C. et al. Regioselective one-pot protection of carbohydrates. Nature446, 896–899 (2007).
Babu, R. S., Chen, Q., Kang, S. W., Zhou, M. & O’Doherty, G. A. De novo asymmetric synthesis of all-d-, all-l-, and d-/l-oligosaccharides using atom-less protecting groups. J. Am. Chem. Soc.134, 11952–11955 (2012).
Koeller, K. M. & Wong, C.-H. Complex carbohydrate synthesis tools for glycobiologists: enzyme-based approach and programmable one-pot strategies. Glycobiology10, 1157–1169 (2000).
Astronomo, R. D. & Burton, D. R. Carbohydrate vaccines: developing sweet solutions to sticky situations?. Nat. Rev. Drug Discov.9, 308–324 (2010).
Vaaje-Kolstad, G. et al. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science330, 219–222 (2010).
Horn, S., Vaaje-Kolstad, G., Westereng, B. & Eijsink, V. G. Novel enzymes for the degradation of cellulose. Biotechnol. Biofuels5, 45 (2012).
Johansen, K. S. Lytic polysaccharide monooxygenases: the microbial power tool for lignocellulose degradation. Trends Plant Sci.21, 926–936 (2016).
Forsberg, Z. et al. Cleavage of cellulose by a CBM33 protein. Protein Sci.20, 1479–1483 (2011).
Quinlan, R. J. et al. Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc. Natl. Acad. Sci. U. S. A.108, 15079–15084 (2011).
Hemsworth, G. R., Johnston, E. M., Davies, G. J. & Walton, P. H. Lytic polysaccharide monooxygenases in biomass conversion. Trends Biotechnol.33, 747–761 (2015).
Harreither, W. et al. Cellobiose dehydrogenase from the ligninolytic basidiomycete Ceriporiopsis subvermispora. Appl. Environ. Microbiol.75, 2750–2757 (2009).
Sygmund, C. et al. Characterization of the two Neurospora crassa cellobiose dehydrogenases and their connection to oxidative cellulose degradation. Appl. Environ. Microbiol.78, 6161–6171 (2012).
Parikka, K. et al. Oxidation of polysaccharides by galactose oxidase. J. Agric. Food Chem.58, 262–271 (2010).
Sygmund, C. et al. Simple and efficient expression of Agaricus meleagris pyranose dehydrogenase in Pichia pastoris. Appl. Microbiol. Biotechnol.93, 695–704 (2012).
Sygmund, C. et al. Characterization of pyranose dehydrogenase from Agaricus meleagris and its application in the C-2 specific conversion of d-galactose. J. Biotechnol.133, 334–342 (2008).
Vuong, T. V. et al. Xylo- and cello-oligosaccharide oxidation by gluco-oligosaccharide oxidase from Sarocladium strictum and variants with reduced substrate inhibition. Biotechnol. Biofuels6, 148 (2013).
Schrewe, M., Julsing, M. K., Bühler, B. & Schmid, A. Whole-cell biocatalysis for selective and productive C–O functional group introduction and modification. Chem. Soc. Rev.42, 6346–6377 (2013).
Xavier, N. M., Rauter, A. P. & Queneau, Y. Carbohydrate-based lactones: synthesis and applications. Top. Curr. Chem.295, 19–62 (2010).
Roduner, E. et al. Selective catalytic oxidation of C-H bonds with molecular oxygen. ChemCatChem.5, 82–112 (2013).
Beeson, W. T., Phillips, C. M., Cate, J. H. D. & Marletta, M. A. Oxidative cleavage of cellulose by fungal copper-dependent polysaccharide monooxygenases. J. Am. Chem. Soc.134, 890–892 (2012).
Agger, J. W. et al. Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation. Proc. Natl. Acad. Sci. U. S. A.111, 6287–6292 (2014).
Couturier, M. et al. Lytic xylan oxidases from wood-decay fungi unlock biomass degradation. Nat. Chem. Biol.14, 306–310 (2018).
Xu, C., Spadiut, O., Araújo, A. C., Nakhai, A. & Brumer, H. Chemo-enzymatic assembly of clickable cellulose surfaces via multivalent polysaccharides. Chemsuschem5, 661–665 (2012).
Aumala, V. et al. Biocatalytic Production of Amino Carbohydrates through Oxidoreductase and Transaminase Cascades. Chemsuschem12, 848–857 (2019).
Isaksen, T. et al. A C4-oxidizing lytic polysaccharide monooxygenase cleaving both cellulose and cello-oligosaccharides. J. Biol. Chem.289, 2632–2642 (2014).
Petrović, D. M. et al. Comparison of three seemingly similar lytic polysaccharide monooxygenases from Neurospora crassa suggests different roles in plant biomass degradation. J. Biol. Chem.294, 15068–15081 (2019).
Vu, V. V., Beeson, W. T., Phillips, C. M., Cate, J. H. D. & Marletta, M. A. Determinants of regioselective hydroxylation in the fungal polysaccharide monooxygenases. J. Am. Chem. Soc.136, 562–565 (2014).
Flitsch, A. et al. Cellulose oxidation and bleaching processes based on recombinant Myriococcum thermophilum cellobiose dehydrogenase. Enzyme Microb. Technol.52, 60–67 (2013).
Westereng, B., Arntzen, M., Agger, J. W., Vaaje-Kolstad, G. & Eijsink, V. G. H. Analyzing activities of lytic polysaccharide monooxygenases by liquid chromatography and mass spectrometry. Methods Mol. Biol.1588, 71–92 (2017).
Westereng, B. et al. Simultaneous analysis of C1 and C4 oxidized oligosaccharides, the products of lytic polysaccharide monooxygenases acting on cellulose. J. Chromatogr. A1445, 46–54 (2016).
MacCormick, B., Vuong, T. V. & Master, E. R. Chemo-enzymatic synthesis of clickable xylo-oligosaccharide monomers from hardwood 4-O-methylglucuronoxylan. Biomacromol19, 521–530 (2018).
Eibinger, M. et al. Cellulose surface degradation by a lytic polysaccharide monooxygenase and its effect on cellulase hydrolytic efficiency. J. Biol. Chem.289, 35929–35938 (2014).
Vuong, T. V., Liu, B., Sandgren, M. & Master, E. R. Microplate-based detection of lytic polysaccharide monooxygenase activity by fluorescence-labeling of insoluble oxidized products. Biomacromol18, 610–616 (2017).
Wang, D. et al. Production of functionalised chitins assisted by fungal lytic polysaccharide monooxygenase. Green Chem.20, 2091–2100 (2018).
Hill, S. & Galan, M. C. Fluorescent carbon dots from mono-and polysaccharides: synthesis, properties and applications. Beilstein J. Org. Chem13, 675–693 (2017).
Leth, M. L. et al. Differential bacterial capture and transport preferences facilitate co-growth on dietary xylan in the human gut. Nat. Microbiol.3, 570–580 (2018).
Hehemann, J. H. et al. Single cell fluorescence imaging of glycan uptake by intestinal bacteria. ISME J.13, 1883–1889 (2019).
Jäger, M. & Minnaard, A. J. Regioselective modification of unprotected glycosides. Chem. Commun.52, 656–664 (2016).
Freimund, S., Huwig, A., Giffhorn, F. & Köpper, S. Rare keto-aldoses from enzymatic oxidation: substrates and oxidation products of pyranose 2-oxidase. Chem. Eur. J.4, 2442–2455 (1998).
Seeberger, P. H. & Werz, D. B. Automated synthesis of oligosaccharides as a basis for drug discovery. Nat. Rev. Drug Discov.4, 751–763 (2005).
Saloheimo, M. et al. EGIII, a new endoglucanase from Trichoderma reesei: the characterization of both gene and enzyme. Gene63, 11–21 (1988).
Vuong, T. V. & Wilson, D. B. The absence of an identifiable single catalytic base residue in Thermobifida fusca exocellulase Cel6B. FEBS J.276, 3837–3845 (2009).
Zatta, P. F. et al. A solid-phase assay for β-1,4-galactosyltransferase activity in human serum using recombinant aequorin. Anal. Biochem.194, 185–191 (1991).
Drueckhammer, D. G. et al. Enzyme catalysis in synthetic carbohydrate chemistry. Synthesis7, 499–525 (1991).
Packer, N. H., Lawson, M. A., Jardine, D. R. & Redmond, J. W. A general approach to desalting oligosaccharides released from glycoproteins. Glycoconj. J.15, 737–747 (1998).
Mravec, J. et al. Tracking developmentally regulated post-synthetic processing of homogalacturonan and chitin using reciprocal oligosaccharide probes. Development141, 4841–4850 (2014).
Westereng, B. et al. Efficient separation of oxidized cello-oligosaccharides generated by cellulose degrading lytic polysaccharide monooxygenases. J. Chromatogr. A1271, 144–152 (2013).
Samuelsen, A. B. et al. Structural features and anti-complementary activity of some heteroxylan polysaccharide fractions from the seeds of Plantago major L. Carbohydr. Polym.38, 133–143 (1999).
Domon, B. & Costello, C. E. A systematic nomenclature for carbohydrate fragmentations in FAB-MS/MS spectra of glycoconjugates. Glycoconj. J.5, 397–409 (1988).
Coenen, G. J., Bakx, E. J., Verhoef, R. P., Schols, H. A. & Voragen, A. G. J. Identification of the connecting linkage between homo- or xylogalacturonan and rhamnogalacturonan type I. Carbohydr. Polym.70, 224–235 (2007).
Westereng, B. et al. Release and characterization of single side chains of white cabbage pectin and their complement-fixing activity. Mol. Nutr. Food Res.53, 780–789 (2009).
Acknowledgements
This work was supported by the Norwegian Research council through grants 244259, 221576, 226244 and 214613, by the Innovation Fund Denmark, Case No: 0603-00522B, and by the Villum VKR Planet project Nr. 00009283.
Author information
Authors and Affiliations
Contributions
B.W., S.K., S.L., M.Ø.A. and F.L.A. designed and performed experiments and analyzed the data. B.W., S.K and V.G.H.E. wrote the main manuscript text. All authors contributed to analysis of the results. All authors reviewed the manuscript. BW and S.K contributed equally to this study.
Corresponding authors
Ethics declarations
Competing interests
The authors declare no competing interests.
Additional information
Publisher's note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Supplementary information
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as 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 images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.
About this article
Cite this article
Westereng, B., Kračun, S.K., Leivers, S. et al. Synthesis of glycoconjugates utilizing the regioselectivity of a lytic polysaccharide monooxygenase. Sci Rep 10, 13197 (2020). https://doi.org/10.1038/s41598-020-69951-7
Received:
Accepted:
Published:
DOI: https://doi.org/10.1038/s41598-020-69951-7
- Springer Nature Limited
This article is cited by
-
Microbial β-glucanases: production, properties, and engineering
World Journal of Microbiology and Biotechnology (2023)
-
Technical pipeline for screening microbial communities as a function of substrate specificity through fluorescent labelling
Communications Biology (2022)