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# In Vitro Synthesis and Crystallization of β-1,4-Mannan

Florent Grimaud, Sandra Pizzut-Serin, Laurence Tarquis, Simon Ladevèze, Sandrine Morel, Jean-Luc Putaux, Gabrielle Potocki-Veronese

► To cite this version:

Florent Grimaud, Sandra Pizzut-Serin, Laurence Tarquis, Simon Ladevèze, Sandrine Morel, et al. In Vitro Synthesis and Crystallization of β-1,4-Mannan. Biomacromolecules, 2019, 20 (2), pp.846-853. 10.1021/acs.biomac.8b01457 . hal-02177345

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# In Vitro Synthesis and Crystallization of β-1,4-Mannan

Florent Grimaud,† Sandra Pizzut-Serin,† Laurence Tarquis,† Simon Ladevèze,† Sandrine Morel,† Jean-Luc Putaux,‡ and Gabrielle Potocki-Veronese*,†

†LISBP, CNRS, INRA, INSAT, Université de Toulouse, F-31400 Toulouse, France

‡Univ. Grenoble Alpes, CNRS, CERMAV, F-38000 Grenoble, France

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## Supporting Information

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**ABSTRACT:** *In vitro* polymerization of *β*-mannans is a challenging reaction due to the steric hindrance conferred by the configuration of mannosyl residues and the thermodynamic instability of the *β*-anomer. Whatever the approach used to date— whether chemical, or enzymatic with glycosynthases and mannosyltransferases—pure *β*-1,4-mannans have never been synthesized *in vitro*. This has limited attempts to investigate their role in the production of plant and algal cell walls, in which they are highly abundant. It has also impeded the exploitation of their properties as biosourced materials. In this paper, we demonstrate that TM1225, a thermoactive glycoside phosphorylase from the hyperthermophile species *Thermotoga maritima*, is a powerful biocatalytic tool for the ecofriendly synthesis of pure *β*-1,4-mannan. The recombinant production of this enzyme and its biochemical characterization allowed us to prove that it catalyzes the reversible phosphorolysis of *β*-1,4-mannosides, and determine its role in the metabolism of the algal mannan on which *T. maritima* feeds in submarine sediments. Furthermore, after optimizing the reaction conditions, we exploited the synthetic ability of TM1225 to produce *β*-1,4-mannan *in vitro*. At 60 °C and from *D*-mannose 1-phosphate and mannohexaose, the enzyme synthesized mannoside chains with a degree of polymerization up to 16, which precipitated into lamellar single crystals. The X-ray powder diffraction and base-plane electron diffraction patterns of the lamellar crystals unambiguously show that the synthesized product belongs to the mannan I family previously observed *in planta* in pure linear mannan, such as those of the ivory nut. The *in vitro* formation of these mannan I crystals is likely determined by the high reaction temperature and the narrow chain length distribution of the insoluble chains.

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## INTRODUCTION

Plant cell wall polysaccharides play a major role in maintaining plant structure and, with regard to certain polymers, are also involved in storing carbohydrates in seeds. Their morphology and supramolecular organization mainly depend on their molecular structure, which is itself conditioned by the amount, efficiency and stability of plant cell wall synthesizing enzymes, and by the respective cellular concentrations of the different substrates required for the synthesis to proceed. These conditions are very difficult to reproduce *in vitro*. This limits our ability to investigate cell wall polysaccharide polymerization and structural organization mechanisms. Yet, this knowledge is essential in order to understand how plant structures are organized at the molecular level, and also fully exploit the functionalities of their individual components as biosourced materials.

Pure linear *β*-1,4-mannans are some of the major components of hemicellulose in the primary cell wall,¹ in which they exist both in crystalline and amorphous forms.²,³

They function either as structural polysaccharides, for example as microfibrils in certain algae,³,⁴ or, in nonleguminous seeds, as storage polysaccharides to feed the growing embryo.² Thanks to their dense crystalline packing and water-insolubility, mannan confer hardness and high resistance to mechanical damage to seeds, even after exposure to water.⁵ Because of these properties, mannan-containing seeds such as ivory nuts⁶ could potentially be used in manufacturing, material production or, such as coconut, to improve the texture of food ingredients.⁷ Short *β*-1,4-mannan chains and mannooligosaccharides are also of interest in the production of food, feed and pharmaceuticals.⁸,⁹ They offer health benefits as prebiotics,¹⁰ are antagonistic agents against cancer cells¹¹ and also exhibit immunomodulatory properties.¹²,¹³ These various applications are encouraging the development of processes for

Received: October 1, 2018

Revised: December 6, 2018

Published: December 6, 2018
the *in vitro* synthesis of β-1,4-mannoside in order to avoid the difficulty of extracting it from plants and separating it from other hemicellulosic constituents.

The *in vitro* synthesis of β-mannan has previously been performed using GDP-mannose as substrate, and crude extracts of recombinant insect¹⁴ or plant¹⁵ cells expressing plant β-mannan synthases. However, the molecular structure of the polysaccharides that were produced in these studies was not characterized. Mannooligosaccharides have also been synthesized using glycosynthases (mutated mannosidases) from α-mannosyl fluoride, a highly unstable chemically modified glycosyl donor¹⁶ (see ref 17 for a review), and also using synthetic chemistry.¹⁸ However, the degree of polymerization (DP) obtained in these enzymatic and chemical approaches was limited to 4 and 8, respectively.

Recently, mannoside phosphorylases have been discovered, as a new class of mannoside-synthesizing enzymes.¹⁹ These bacterial enzymes, which act on β-1,2,²⁰ 1,3²¹ or 1,4-(hetero)mannosides,¹⁹,²² are classified in the CAZy database as belonging to the GH130 family,²³ which today contains more than 1500 members, of which only 16 have been functionally characterized. The physiological role of mannoside phosphorylases is the degradation of mannosides by phosphorolysis. This reaction being reversible, these enzymes can synthesize β-mannosides *in vitro* from α-D-mannose 1-phosphate (αMan1P) and various acceptors. Therefore, they represent an attractive prospect as green catalysts for the selective synthesis of β-mannosides with a specific linkage composition and high level of purity. The mannoside phosphorylase that has been found to synthesize the longest β-1,4-mannooligosaccharides (DP 12) *in vitro* is UhgbMP, a rather unstable enzyme produced by an uncultured human gut bacterium in order to degrade human N-glycans.²² One of its homologues, the TM1225 protein from *Thermotoga maritima* MSB8, which shares a 61% identity with UhgbMP, was the first GH130 member to be structurally characterized. Although its crystallographic structure (PDB accession number 1VKD) was established in 2004, this protein has still not been biochemically characterized. In the *Thermotoga maritima* MSB8 genome,²⁴ the TM1225 encoding gene belongs to one of the two β-mannan utilization loci.²⁵⁻²⁷ *Thermotoga maritima* MSB8 is a hyperthermophile bacterium, isolated from sediments collected near a submarine hot spring in the Bay of Naples,²⁸ which feeds on several plant and algal cell wall constituents, such as β-mannans.²⁶,²⁹

In this study, we characterized the function of TM1225 in order to first determine what role it plays in the mannan metabolism of *Thermotoga maritima* MSB8. After optimizing the reaction conditions, we exploited its high thermoactivity and stability to synthesis mannan by reverse phosphorolysis. *In vitro*, these pure linear β-1,4-mannan chains assemble into crystals, whose morphology and structure were characterized in detail and compared to those observed either *in planta* or after crystallizing native mannans *in vitro*.

## ■ MATERIAL AND METHODS

**Production and Purification of the Recombinant TM1225 Enzyme.** The gene encoding the TM1225 protein from *Thermotoga maritima* strain ATCC 43589/MSB8/DSM 3109/JCM 10099 (TM1225, Genbank accession number AAD36300.1, PDB 1VKD) was synthesized and cloned into the pET28a vector by the Biomatik Company (Biomatik, Ontario, Canada) between restriction sites NcoI and XhoI. The resulting plasmid sequence is provided in the Supporting Information. The recombinant enzyme was produced as

the fusion protein (His)₆-TM1225 by *Escherichia coli* BL21 star cells carrying the pET28a-tm1225 plasmid. For the sake of simplicity, (His)₆-TM1225 will hereafter be referred to as TM1225. The culture was performed for 24 h at 20 °C in the ZYM-50S2 autoinductible medium supplemented with 50 µg mL⁻¹ kanamycin, and inoculated at A₆₀₀ nm 0.1. The cells were harvested and resuspended in 20 mM Tris-HCl at pH 7.0 and 300 mM NaCl, and lysed using sonication. The soluble lysate was applied to a TALON resin loaded with cobalt (GE Healthcare) equilibrated in 20 mM Tris-HCl at pH 7.0 and 300 mM NaCl. After column washing with eight volumes of this same buffer supplemented with 10 mM imidazole, the protein was eluted in 20 mM Tris-HCl at pH 7.0, 300 mM NaCl and 250 mM imidazole. Finally, TM1225 was desalted using a gel filtration column (PD10, Amersham), with a 20 mM MES buffer at pH 6.0 as the eluent. The purity of TM1225 was evaluated by SDS-PAGE using Mini PROTEAN TGX Stain-Free Gels (Bio-Rad). Protein concentrations were determined through spectrometry using a NanoDrop ND-1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA). The extinction coefficient of the purified TM1225 fused to the N-terminal His₆ tag was 75145 mol⁻¹ cm⁻¹, a value estimated with the Protparam software (https://web.expasy.org/protparam/).

**In Vitro Phosphorolysis and Reverse Phosphorolysis.** In order to determine TM1225 specificity, the phosphorolysis and reverse phosphorolysis reactions were performed during 24 h at 37 °C in 50 mM Tris-HCl buffer at pH 7.0, using 1.0 µg mL⁻¹ of purified TM1225. Phosphorolysis was performed in the presence of 10 mM inorganic phosphate and 10 mM 1,4-β-D-mannooligosaccharides as substrates (mannobiose (M2), mannotriose (M3), mannotetraose (M4) and mannopentaose (M5) from Megazyme, Ireland). Reverse phosphorolysis was performed using αMan1P (sodium salt hydrate, Sigma-Aldrich, reference M1755) at 10 mM as the mannosyl donor, with 10 mM D-mannose as acceptor. The best conditions for reverse phosphorolysis were determined by incubating during 24 h 1.0 µg mL⁻¹ of purified TM1225 at different temperatures (from 30 to 80 °C) and pH levels (from 4.0 to 9.0), with 10 mM αMan1P as the mannosyl donor and 10 mM D-mannose as the acceptor. To determine the optimal pH, the assays were performed at 37 °C using a 20 mM Tris-HCl buffer (for pH values of between 3.5 and 5.5), 20 mM MES buffer (for pH values between 6.0 and 6.5) and 20 mM sodium acetate buffer (for pH values between 7.0 and 9.0). The optimal temperature was then determined at the optimal pH (pH 6.0). All these reactions were performed in a final volume of 500 µL. The reactions were stopped by sample incubation at 95 °C during 5 min. We checked by HPAEC-PAD analysis of the reaction medium that TM1225 was not active anymore after this treatment. Substrate and product concentrations were determined by HPAEC-PAD analysis, as further described in this section. The percentage of mannosyl groups incorporated into β-1,4-D-mannoside chains from D-mannose and αMan1P in the optimal conditions of reaction was calculated as follows:

$$ \%_{\text{Mannosyl groups incorporated into mannosides}} = \frac{\sum_i [\text{Ml}]_{\text{final}}}{[\alpha\text{Man1P}]_{\text{initial}} + [\text{D-Mannose}]_{\text{initial}}} \times 100 $$

where [Ml]final is the final molar concentration of β-1,4-D-mannosides with a DP ranging from 2 to 6, [αMan1P]initial is the initial molar concentration of αMan1P (10 mM) and [D-Mannose]initial is the initial molar concentration of D-mannose (10 mM).

Mannan synthesis from 10 mM αMan1P and 10 mM mannohexaose was performed during 24 h at 60 °C, using 1.0 µg mL⁻¹ of purified TM1225 in a 20 mM MES buffer at pH 6.0, in a final reaction volume of 10 mL. In this case, the reaction was not stopped by heating, in order not to affect mannan precipitation and crystallization. After reaction and centrifugation of the reaction medium at 10,000g for 10 min, the products were immediately analyzed by HPAEC-PAD analysis as further described in this section. The reaction precipitate was washed 3 times in one volume of 20 mM MES at pH 6.0 in order to eliminate the enzyme, and finally resuspended in the same amount of buffer as in the initial reaction
medium for transmission electron microscopy and X-ray diffraction analyses.

**Characterization of the Structure of the TM1225 Products.**
*High-Performance Anion-Exchange Chromatography with Pulsed Amperometric Detection (HPAEC-PAD)*. The chain length distribution of the reaction products was determined using HPAEC-PAD. Aliquots of the reaction media were centrifuged at 10,000g for 10 min, and the soluble fractions (supernatants) were diluted 100 times with water. For reactions in which an insoluble fraction was obtained (i.e., TM1225 with αMan1P and mannohexaose), the reaction media were first centrifuged at 10,000g for 10 min and divided into two fractions: the soluble fraction (supernatant) and the insoluble fraction (precipitate). The insoluble fraction was washed 3 times in one volume of 20 mM MES at pH 6.0, and finally resuspended in the same amount of buffer as in the initial reaction medium. Finally, all fractions were solubilized using NaOH 0.1 M final (100 µL of the sample was solubilized by the addition of 2 µL of 5 M NaOH). An extemporary analysis was performed on the resulting fractions using HPAEC-PAD to determine the mannoside chain length distribution. The HPAEC-PAD analysis was performed on a 2 × 250 mm Dionex Carbopac PA100 column. A gradient of sodium acetate in 150 mM NaOH at a 0.25 mL min⁻¹ flow rate was applied as follows: 0–15 min, 0–150 mM; 15–18 min, 140–140 mM; 18–20 min, 140–0 mM; 20–25 min, 0–0 mM. The detection was performed using a Dionex ED40 module with a gold working electrode and an Ag/AgCl pH reference.

*¹H and ¹³C NMR*. The freeze-dried reaction media was resuspended in deuterated water. Deuterium oxide was used as the solvent, and sodium 2,2,3,3-tetradeu terio-3-trimethylsilylpropanoate was selected as the internal standard. ¹H and ¹³C NMR spectra were recorded on an Advance 500-MHz spectrometer (Bruker) operating at 500.13 MHz for ¹H NMR and 125.75 MHz for ¹³C using a 5 mm z-gradient TBI probe. The data have been acquired at 298 K, and processed using TopSpin 3.0 software. The various signals were assigned by comparison with those obtained from standards (αMan1P and 4-O-β-D-mannopyranosyl-D-mannose) as described.²²

*Transmission Electron Microscopy (TEM)*. Drops of 10 µg mL⁻¹ carbohydrate suspensions were deposited onto glow-discharged carbon-coated grids and negatively stained with 20 mg mL⁻¹ uranyl acetate. The preparations were observed using a FEI-Philips CM200 transmission electron microscope operating at 200 kV. Unstained specimens where probed at room temperature using selected area electron diffraction (ED). The images and ED patterns were recorded using a TVIPS TemCam F216 camera.

*X-ray Diffraction (XRD)*. The insoluble fraction was air-dried and the resulting powder was poured into a 1 mm (outer diameter) glass capillary that was flame-sealed. The specimen was X-rayed in a vacuum by Ni-filtered Cu Kα radiation (λ = 0.154 18 nm) using a Philips PW3830 generator operating at 30 kV and 20 mA. A two-dimensional diagram was recorded on a Fujifilm image plate and read using a Fujifilm BAS 1800-II bioimaging analyzer. The XRD profile was calculated by rotationally averaging the 2D pattern.

## RESULTS AND DISCUSSION

**Function of TM1225 in Thermotoga maritima.** The mannan catabolic machinery of *Thermotoga maritima* MSB8 has previously been partly characterized.³,³¹ It has been proposed that the TM1225 protein catalyzes the intracellular conversion of *β*-1,4-*D*-mannosides, the result of the hydrolysis of high molecular mass mannans by the extracellular (TM1227) and intracellular *β*-mannanases (TM1751 and TM1752), into *D*-mannose and mannose 6-phosphate.³⁰ However, TM1225 is classified as belonging to the GH130 family, which contains several *β*-mannoside phosphorylases able to break down *β*-mannoside into *α*Man1P and *β*-mannosides of lower chain length. We thus speculated that the physiological role of TM1225 would be the intracellular conversion of *β*-1,4-*D*-mannosides into *α*Man1P and, ultimately, *D*-mannose. In order to characterize the GH130

enzyme TM1225, and to confirm its function in *Thermotoga maritima*, we produced it in *E. coli*, in fusion with an N-terminal (His)6 tag, according to the protein production process that led to the resolution of the TM1225 crystallographic structure (1VKD). The recombinant enzyme was purified to homogeneity using affinity chromatography (Figure 1).

<figure><img src="image_7.png" /><figcaption>**Figure 1.** Purification of the mannoside phosphorylase TM1225. Protein fractions were separated by SDS-PAGE with Stain-Free Precast Gels that allow rapid fluorescent detection of proteins. Lane S, molecular weight standards; lane 1, cell sonication supernatant; lane 2, purification flowthrough; lane 3, purified TM 1225 in the elution fraction; lane 4, purified TM 1225 after desalting.</figcaption></figure>

Incubations of TM1225 in the presence of inorganic phosphate and *β*-1,4-*D*-mannosides with a DP ranging from 2 to 5 (M2 to M5) resulted in the conversion of the substrates into mannosides with a size either larger or smaller than that of the initial chains, as well as a product of which the elution time was strictly equivalent to that of *α*Man1P, which is very different from that *D*-mannose 6-phosphate (Man6P) (Figure 2).

On the basis of these findings, we evaluated the ability of TM1225 to perform the reverse phosphorolysis reaction using *α*Man1P (10 mM) as the mannosyl donor and *D*-mannose (10 mM) as the acceptor (Figure 3A).

The reaction resulted in the production of oligosaccharides with retention times corresponding to those of standard *β*-1,4-*D*-mannosides with a DP ranging from 1 to 5 (M1 to M5). ¹H and ¹³C NMR analyses of these products, confirmed the ability of TM1225 to catalyze the formation of *β*-1,4-*D*-linked mannopyranosyl residues (Figure 3B,C). No other product than *D*-mannose, *β*-1,4-*D*-mannosides and *α*Man1P was observed on the NMR spectra. Taken together, these results confirm that TM1225 is involved in the reversible phosphorolysis of *β*-1,4-*D*-mannosides as follows: (β-1,4-D-mannosides)ₙ + Pi ⇌ (β-1,4-D-mannosides)ₙ-₁ + αMan1P. The final phosphorolysis products of TM1225 (*α*Man1P and *D*-mannose) could serve as key intermediates for energy production and glycoprotein formation by *Thermotoga maritima*, as previously described for other GH130 enzymes.²⁰,³² Like for other anaerobes, using phosphorolysis
<figure><img src="image_8.png" /><figcaption>**Figure 2.** Phosphorolysis reactions catalyzed by TM1225. HPAEC-PAD chromatograms of the reaction products obtained after incubation of TM1225 with 1,4-β-D-mannosides with a DP ranging from 2 to 5 (M2 to M5) (10 mM) and inorganic phosphate (10 mM). HPAEC-PAD chromatograms of αMan1P and Man6P standards are also represented.</figcaption></figure>

<figure><img src="image_9.png" /><figcaption>**Figure 3.** Reverse phosphorolysis catalyzed by TM1225. (A) HPAEC-PAD chromatograms of the reaction products obtained after incubation of TM1225 with D-mannose (10 mM) and αMan1P (10 mM). ¹H (B) and ¹³C (C) NMR spectra of the products synthesized during reverse phosphorolysis.</figcaption></figure>

rather than hydrolysis may be helpful for its growth, phosphorolysis saving ATP compared to hydrolysis and monosaccharide activation to sugar-phosphates.<sup>33,34</sup>

**β-1,4-D-Mannan Synthesis by Reverse Phosphorolysis.** The second objective of this work was to use TM1225 as a biotechnological tool for the synthesis of β-1,4-D-mannan from αMan1P as the mannosyl donor, and D-mannose or β-1,4-D-mannosides as acceptors. We therefore looked at the impact of the reaction physicochemical parameters and acceptor chain length on the yield of the reverse phosphorolysis products. First, the pure enzyme was incubated with D-mannose (10

mM) and αMan1P (10 mM) at different temperatures and pH levels (from 30 to 80 °C and from 4.0 to 9.0, respectively). The reaction products were quantified using HPAEC-PAD (Figure 4). In all cases, the reactions resulted in the production of a series of β-1,4-D-mannosides with a DP ranging from 1 to 6. The highest concentration of β-1,4-D-mannosides was obtained at 60 °C and pH 6.0. In these conditions, the percentage of mannosyl groups incorporated from D-mannose and αMan1P into β-1,4-D-mannoside chains was 64.5%.

These conditions were thus considered to be the optimal conditions for β-1,4-D-mannoside synthesis by reverse
<figure><img src="image_10.png" /><figcaption>**Figure 4.** Determination of the optimal temperature and pH for β-1,4-D-mannoside synthesis by reverse phosphorolysis. Quantification using HPAEC-PAD of the reaction products obtained from D-mannose (10 mM) and αMan1P (10 mM).</figcaption></figure>

<figure><img src="image_11.png" /><figcaption>**Figure 5.** Reverse phosphorolysis from αMan1P and M1 or M6. HPAEC-PAD analysis of the TM1225 reaction products obtained in the presence of 10 mM M1 or M6, and 10 mM αMan1P. The reaction precipitate was solubilized in 0.1 M NaOH before HPLC analysis. No precipitate was obtained in the presence of M1 as the acceptor.</figcaption></figure>

phosphorolysis. In order to orient the reverse phosphorolysis reaction toward the synthesis of β-1,4-D-mannosides with higher DPs, TM1225 was then incubated with 10 mM mannohexose (M6) and αMan1P. The formation of β-1,4-D-mannosides was tracked using HPAEC-PAD (Figure 5).

Under these conditions, TM1225 performed two different reactions: (i) mannohexaose phosphorolysis producing β-1,4-D-mannosides with lower DPs (M5 to M1), and (ii) the synthesis of soluble β-1,4-D-mannosides with higher DPs (M7 to M11) by reverse phosphorolysis. In addition, a precipitate was formed during the course of the reaction, a phenomenon which was not observed when D-mannose was used as the acceptor.

**Structural Characterization of the Reaction Precipitate.** In order to determine the nature and composition of the precipitate produced from 10 mM M6 and 10 mM αMan1P, the sample was fractionated by centrifugation, and the precipitate was solubilized in NaOH before the HPAEC-

PAD analysis (Figure 5). As previously explained, the soluble fraction contained a mixture of β-1,4-D-mannoside chains ranging from DP 1 to 11. The total fraction contained M1 to M16, with a higher amount of β-1,4-D-mannoside from DP 8 to 16 compared to the soluble fraction. After solubilization of the reaction precipitate in NaOH, we observed on the HPAEC-PAD chromatogram that it mainly contained β-1,4-D-mannosides ranging from M7 to M16 (Figure 5), indicating that the DP 7–11 chains partially precipitated with the insoluble DP 12–16 products during the reaction.

We characterized the structure of the insoluble products using X-ray diffraction. As shown in Figure 6, diffraction peaks at 2θ = 15.9, 18.2, 19.9, 23.5, 25 and 26.8° were detected, indicating that the β-1,4-D-mannoside chains crystallized during reverse phosphorolysis. The diffraction pattern is identical to those of mannan I produced by recrystallizing short mannan chains from ivory nuts.³⁵,³⁶
<figure><img src="image_12.png" /><figcaption>**Figure 6.** (A) XRD powder pattern of the insoluble fraction; (B) corresponding XRD profile.</figcaption></figure>

TEM images of negatively stained preparations clearly show the presence of stacks of parallelepipedal lamellar crystals (Figure 7A). The lozenge shape is clearer for isolated stacks

<figure><img src="image_13.png" /><figcaption>**Figure 7.** (A) TEM image of lamellar crystals in the insoluble fraction formed in the TM1225 reaction media, negatively stained with uranyl acetate; (B) TEM image of unstained crystals; (C) base-plane electron diffraction pattern recorded at low temperature and properly oriented with respect to image B.</figcaption></figure>

such as that shown in Figure 7B. Unstained single crystals yielded a base-plane ED pattern identical to that found for crystals prepared by recrystallizing ivory nut mannans,³⁵,³⁶ which confirms the mannan I nature of the crystals.

Previous crystallization studies with ivory nut linear mannan and glucomannan indicated that, depending on parameters like crystallization temperature, medium polarity and chain length distribution, two distinct allomorphs could be obtained.³⁷⁻⁴⁰ In mannan I, antiparallel chains are tightly packed in an orthorhombic unit cell, and the crystal does not contain any intracrystalline water molecule.³⁶,³⁹,⁴⁰ The structure of mannan II is not as well-known and its antiparallel chains would be

associated with a number of water molecules.³⁸,⁴¹,⁴² In a previous study, a high temperature, low polarity and short chain length have been found to favor the formation of mannan I.³⁷ In this study, the *in vitro* synthesis was performed at a high temperature and resulted in the formation of short β-1,4-D-mannoside chains (1 < DP < 16) that spontaneously crystallized without adding any exogeneous precipitant to the reaction medium. The conditions used here for reverse phosphorolysis were thus suitable for the formation of mannan I, one of the allomorphs produced *in planta* with mannan chains with a DP of around 15.

*In planta*, mannan has been found with various morphologies (microfibrils, ribbons, granules) and allomorphic structures (I and II).³ Mannan I is generally granular, while mannan II is microfibrillar in ivory nuts, the cell wall of some algae and the endosperm of some Leguminosae.³,⁵,⁴³ The crystal morphology drastically changes after the dissolution and recrystallization of mannan fractions. The low-DP chains can form lamellar crystals with a lozenge shape³⁵ or ribbons.³⁷,⁴⁴ Heux et al.⁴² have reported the formation of ribbon-like mannan II by crystallizing DP 30 chains from ivory nuts and a DP 800 fraction from *Acetabularia crenulata*.

Crystallization during *in vitro* enzymatic polymerization and without the addition of a precipitating agent has been reported for β-1,4-⁴⁵⁻⁴⁷ and β-1,3-D-glucans,⁴⁸ as well as for those containing a mixture of both types of linkages.⁴⁹ The glucan chains can precipitate in the form of spherulitic particles composed of crystallites organized radially,⁵⁰ or directly as lamellar single crystals.⁴⁸ Nevertheless, the phenomena involved in the *in vitro* crystallization of these glucans and mannan cannot be compared, since the reaction conditions (synthesizing enzymes, temperature, substrate concentrations) and polymer structures (glycosyl type and osidic linkage) are very different. Glycoside-phosphorylases, some of which have been found to be efficient polymerases in recent years, will be interesting tools for diversifying the examples of polysaccharide synthesis *in vitro*, and for understanding the relationships between molecular structure and crystallization properties, by varying only one or several reaction parameters.

<figure><img src="image_14.png" /></figure>

## CONCLUSION

*Thermotoga maritima*, a thermophilic bacterial species that feeds on numerous plant polysaccharides, is a goldmine in the search for thermostable enzymes of biotechnological interest. In this study, we characterized the function of TM1225, which became, in 2004, the first GH130 member to have its 3D structure released in the Protein Data Bank, although its function was still a matter of speculation. Here, we have demonstrated that TM1225 is a β-1,4-D-mannoside phosphor-ylase able to act on chains with a DP ≥ 2 according to our previous prediction based on structural comparison analyses,⁵¹ but not with other hypotheses regarding its physiological role.³⁰ We further demonstrated the ability of TM1225 to synthesize *in vitro* β-1,4-D-mannoside chains up to DP 16, a length that had never been reached through chemical¹²,¹⁸,⁵²,⁵³ or enzymatic synthesis using glycosynthases.¹⁷ The thermoac-tive TM1225 mannoside phosphorylase, which can easily be produced in *E. coli* and purified, thus represents the most interesting enzymatic tool to date for the one-step synthesis of mannosides. The high temperature used for reverse phosphor-olysis and the narrow chain length distribution of the mannoside chains are probably the major determinants of
their precipitation *in vitro* into mannan I crystals, like those produced *in planta*.

<figure><img src="image_15.png" /></figure>

## ASSOCIATED CONTENT

<figure><img src="image_16.png" /></figure>

### Supporting Information

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.8b01457.

Sequence of the Pet28a-tm1225 plasmid (PDF)

<figure><img src="image_17.png" /></figure>

## AUTHOR INFORMATION

### Corresponding Author

*Gabrielle Potocki-Veronese. E-mail: veronese@insa-toulouse.fr.

### ORCID

<figure><img src="image_18.png" /></figure>

Florent Grimaud: 0000-0003-0614-5211

### Notes

The authors declare no competing financial interest.

<figure><img src="image_19.png" /></figure>

## ACKNOWLEDGMENTS

The HPLC analyses were performed using the ICEO facility, which is dedicated to the screening, discovery and characterization of new enzymes. ICEO, part of the Integrated Screening Platform of Toulouse (PICT, IBiSA), is supported by grants from the Région Midi-Pyrénées, the European Regional Development Fund and the Institut National de la Recherche Agronomique (INRA). We are grateful to MetaSys, the Metabolomics and Fluxomics Center at the Laboratory for the Engineering of Biological Systems and Processes (Toulouse, France) for the NMR experiments, and to the NanoBio-ICMG Platform (FR 2607, Grenoble), for granting access to the Electron Microscopy facility. We cordially thank Nelly Monties and Angeline Rizzo for their technical support. This work was financed by the Agence Nationale de la Recherche (project ASPIRE, grant number ANR-14-CE27-0011-02, and project Oligomet, grant number ANR-16-CE20-0006) and by the European Union's framework programme Horizon 2020 (LEIT-BIO-2015-685474, Metafluidics).

<figure><img src="image_20.png" /></figure>

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