A Single Heterologously Expressed Plant Cellulose Synthase Isoform Is Sufficient for Cellulose Microfibril Formation in Vitro

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A single heterologously expressed plant cellulose synthase isoform is sufficient for cellulose microfibril formation in vitro

Pallinti Purushothama, Sung Hyun Chob, Sara M. Díaz-Morenoc, Manish Kumard, B. Tracy Nixonb, Vincent Bulonec,e, and Jochen Zimmera,1

aDepartment of Molecular Physiology and Biological Physics, University of Virginia School of Medicine, Charlottesville, VA 22908; bDepartment of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park, PA 16802; cDivision of Glycoscience, Royal Institute of Technology, Stockholm, SE-10691, Sweden; dDepartment of Chemical Engineering, The Pennsylvania State University, University Park, PA 16802; and eAustralian Research Council Centre of Excellence in Plant Cell Walls, School of Agriculture, Food and Wine, University of Adelaide, Waite Campus, Urrbrae, SA, 5064, Australia

Edited by Chris R. Somerville, University of California, Berkeley, CA, and approved August 9, 2016 (received for review April 18, 2016) Plant cell walls are a composite material of polysaccharides, proteins, In most plant tissues, cellulose polymers are bundled into and other noncarbohydrate polymers. In the majority of plant tissues, cable-like structures that are wrapped around the cell to form the the most abundant polysaccharide is cellulose, a linear polymer of load-bearing component of the cell wall (1). The smallest repeating glucose molecules. As the load-bearing component of the cell wall, unit, the cellulose microfibril, consists of a currently unknown [but individual cellulose chains are frequently bundled into micro and most likely 18–24 (4, 5)] number of cellulose polymers. The mi- macrofibrils and are wrapped around the cell. Cellulose is synthe- crofibrils are stabilized by van der Waals interactions between the sized by membrane-integrated and processive glycosyltransferases polymer’s glucopyranose rings and by hydrophilic intermolecular that polymerize UDP-activated glucose and secrete the nascent contacts mediated by the sugars’ equatorial hydroxyl groups. polymer through a channel formed by their own transmembrane The mechanism by which CesA synthesizes and secretes individual regions. Plants express several different cellulose synthase isoforms glucan chains has been delineated recently for bacterial cellu- during primary and secondary cell wall formation; however, so far, lose synthase (6–8); however, the process by which plant CesAs none has been functionally reconstituted in vitro for detailed bundle these glucan chains into cellulose microfibrils is currently biochemical analyses. Here we report the heterologous expression, unknown. An intriguing hypothesis is that the oligomeric state purification, and functional reconstitution of Populus tremula x of CesAs in the plasma membrane dictates the spontaneous tremuloides CesA8 (PttCesA8), implicated in secondary cell wall association of the extruded glucan chains into microfibrils formation. The recombinant enzyme polymerizes UDP-activated (9, 10). Indeed, plant cellulose synthases have been shown to glucose to cellulose, as determined by enzyme degradation, perme- form large membrane-embedded complexes displaying pseudo-sixfold thylation glycosyl linkage analysis, electron microscopy, and muta- symmetry referred to as “CesA rosettes” (11, 12). In some species genesis studies. Catalytic activity is dependent on the presence of a these rosettes are organized further into 2D arrays from which cel- lipid bilayer environment and divalent manganese cations. Further, lulose fibrils originate (13). Thus it seems likely that close proximity electron microscopy analyses reveal that PttCesA8 produces cellulose of CesAs during cellulose biosynthesis is sufficient for microfibril fibers several micrometers long that occasionally are capped by glob- formation. This notion is supported by the observation that ular particles, likely representing PttCesA8 complexes. Deletion of cellulose produced by monomeric bacterial cellulose synthase is the enzyme’s N-terminal RING-finger domain almost completely amorphous, with no detectable higher-order structure (14). abolishes fiber formation but not cellulose biosynthetic activity. Our results demonstrate that reconstituted PttCesA8 is not only sufficient Significance for cellulose biosynthesis in vitro but also suffices to bundle individual glucan chains into cellulose microfibrils. Cellulose is an abundant natural polymer synthesized primarily by vascular plants in which it forms the load-bearing compo- biopolymer | cellulose | glycosyltransferase | plant cell wall | nent of the cell wall. It is a linear polymer of glucose molecules membrane transport synthesized by membrane-embedded cellulose synthases that couple polymer synthesis with its secretion across the plasma ellulose is an abundant biopolymer primarily produced by membrane. Plants express multiple cellulose synthase isoforms Cvascular plants and also by some bacteria, algae, and tunicates that are organized into large macromolecular assemblies of (1). It is a polymer of glucose molecules connected between their varying composition that are likely responsible for aligning the C1 and C4 carbons via β-glycosidic linkages. The unbranched cellulose strands into microfibrils. Here we show that recom- polymer is further stabilized by intramolecular hydrogen bonds binantly expressed and purified Populus tremula x tremuloides between the ring oxygen and C2 hydroxyl of one sugar and the (hybrid aspen) cellulose synthase-8 is sufficient for cellulose C3 and C6 hydroxyl groups, respectively, of a neighboring biosynthesis and produces cellulose microfibrils in vitro. Our sugar unit. results demonstrate that no other plant-derived factors are Cellulose is synthesized by cellulose synthase, CesA (1), a required for cellulose microfibril biosynthesis. membrane-integrated processive family-2 glycosyltransferase that shares significant similarity with chitin, hyaluronan, and alginate Author contributions: P.P. and J.Z. designed all biochemical experiments; S.H.C., M.K., and B.T.N. designed all EM experiments; S.M.D.-M. and V.B. designed the linkage analysis exper- synthases (2). CesA combines two functions: the enzyme poly- iments; P.P., S.H.C., and S.M.D.-M. performed all experiments; P.P., S.H.C., S.M.D.-M., M.K., merizes UDP-activated glucose (UDP-Glc) in an SN2-like nu- B.T.N., V.B., and J.Z. contributed to analyzing and writing the manuscript. cleophilic displacement reaction (3) and translocates the growing The authors declare no conflict of interest. cellulose polymer across the plasma membrane through a pore This article is a PNAS Direct Submission. formed by its own transmembrane (TM) region. Depending on 1To whom correspondence should be addressed. Email: [email protected]. the tissue, the synthesized polymers can be of astonishing lengths, This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. some containing tens of thousands of sugar units (1). 1073/pnas.1606210113/-/DCSupplemental.

11360–11365 | PNAS | October 4, 2016 | vol. 113 | no. 40 www.pnas.org/cgi/doi/10.1073/pnas.1606210113 Downloaded by guest on September 27, 2021 revealed that the recombinant protein is membrane-integrated, a prerequisite for proper PttCesA8 folding and function (Fig. 1A). Approximately 50% of the enzyme can be solubilized from the membrane in several nondenaturing detergents, including Triton X-100, dodecyl-maltoside (DDM), sodium cholate, and polyoxyethylene(8)dodecyl ether (C12E8). For all subsequent PttCesA8 purifications, DDM was used for solubilization and LysoFoscholine-Ether14 for purification (see Methods). How- ever, membrane isolation and solubilization generated a smaller PttCesA8 species migrating at ∼60 kDa. This species was detectable only with an antibody directed against the C-terminal His-tag and not against the N-terminal FLAG-tag, suggesting that the N-terminal region of PttCesA8 is prone to degradation, likely because of conformational flexibility or loose association of the N-terminal RING-finger (Fig. 1B). Interestingly, cobalt- affinity chromatography primarily enriched the full-length PttCesA8 protein, perhaps because of weaker interactions with the affinity matrix or aggregation of the truncated species (Fig. 1B). Although SDS/PAGE of the affinity-purified fraction followed by Fig. 1. Expression and purification of recombinant PttCesA8. (A, Left) Coomassie and silver staining revealed the presence of only one Western blot analysis of C-terminally His-tagged PttCesA8 expressed in major protein species (Fig. 1B), we determined its total protein P. pastoris. IMV, inverted membrane vesicles; S, soluble fraction; WCE, whole- composition (arising from contaminating Pichia proteins) by cell extract. (Right) Solubilization of IMVs with different nondenaturing detergents. Δ tandem mass spectrometry fingerprinting (Table S1). Apart from C, C12E8; D, DDM; NaC, sodium cholate; T, Triton X-100. FL and N: Full- a small number of peptides matching a Pichia β-1,3 glucan syn- length and truncated CesA8, respectively. (B) Cobalt-affinity purification of PttCesA8. (Left) Western blot against the N-terminal FLAG-tag of PttCesA8. thase, none of the contaminating proteins has any documented (Right) Coomassie (Coo)- and silver (Ag)-stained SDS/PAGE of the eluted or predicted function in polysaccharide biosynthesis. PttCesA8 was fraction. E, eluted fraction; FT, flow-through; L, load; W1–4, wash steps 1–4 represented by 61 unique peptides covering most of the enzyme’s with imidazole concentrations increasing from 20–60 mM. hydrophilic regions.

Recombinant PttCesA8 Synthesizes Cellulose. Following previous Despite many years of study, cellulose biosynthesis has been successes with bacterial cellulose synthase (14), we reconstituted reconstituted in vitro only recently from purified components PttCesA8 into Saccharomyces cerevisiae total lipid extract proteoli- from Rhodobacter sphaeroides (14). In these studies, the cellulose posomes and analyzed the enzyme’s catalytic activity in the presence + synthase was BcsA, which requires a membrane-anchored, periplasmic of UDP-Glc, Mn2 ,andUDP-[3H]-Glc as tracer. Because cellulose subunit (BcsB) for catalytic activity. Plant cellulose biosynthetic activity is water insoluble past a degree of polymerization of about 8 (18), has been demonstrated in membrane extracts from several plant insoluble reaction products were sedimented and further purified tissues (15–17), but none of the catalytically active synthases has by descending paper chromatography as described for bacterial been purified to date. Because of these difficulties, it was spec- ulated that the plant enzymes also might require an additional, weakly associated factor for catalytic activity, similar to the BcsA–B complex in bacteria. Plants produce several CesA isoforms; some are required for

primary and others for secondary cell wall formation (1). To de- PLANT BIOLOGY termine whether a single CesA isoform is catalytically active, we heterologously expressed and purified Populus tremula × tremuloides (hybrid aspen) CesA isoform 8 (PttCesA8), which is implicated in secondary cell wall formation, in Pichia pastoris. The enzyme was reconstituted into proteoliposomes where it synthesizes cellulose microfibrils in an UDP-Glc– and manganese-dependent manner, thereby demonstrating that a single CesA isoform suffices for cellulose biosynthesis and microfibril formation. Results Heterologous Expression and Purification of PttCesA8. CesA is an integral membrane protein predicted to contain an N-terminal cytosolic domain preceding two TM helices, an intracellular glycosyltransferase domain, and five C-terminal TM helices (6). If this prediction is correct, the odd number of TM helices would place the enzyme’s N and C termini on opposing sides of the Fig. 2. In vitro cellulose biosynthesis by reconstituted PttCesA8. (A) Time plasma membrane. To facilitate detection and purification of the course of cellulose biosynthesis using UDP-[3H]-glucose as tracer. DPM, disinte- recombinant enzyme, we engineered an N-terminal FLAG and grations per minute. (B and C) pH and temperature (B) and cation dependence C-terminal His-tag on PttCesA8 (Fig. S1). The construct was (C)ofPttCesA8’s catalytic activity. Synthesis reactions were performed in the expressed in P. pastoris upon induction with methanol, leading to presence of 20 mM of the indicated divalent cations. A catalytically inactive PttCesA8 mutant (CesA8*) and material obtained from a mock purification the formation of an immunoreactive protein species migrating do not produce detectable glucan amounts. (D) Enzymatic degradation of the near the 100-kDa marker on SDS/PAGE, consistent with the PttCesA8-synthesized glucan. All experiments were performed at least in calculated PttCesA8 molecular mass of 110.5 kDa (Fig. 1A). Frac- triplicate; error bars represent the SDs from the means. AG, amyloglucosidase; tionation of the cell lysate into soluble and membrane fractions AL, amylase; GH, glycoside hydrolase.

Purushotham et al. PNAS | October 4, 2016 | vol. 113 | no. 40 | 11361 Downloaded by guest on September 27, 2021 PttCesA8 activity (Fig. S3). In this case, glycosyl linkage analysis of the product also revealed the presence of a 1,3-linked glucan, + most likely arising from a contaminating Mg2 -dependent β-1,3- glucan synthase and not from PttCesA8. As expected for a con- tamination, the presence of the β-1,3-glucan synthase activity varied among purifications, and we used only enzyme preparations exhibiting negligible catalytic activity in the presence of magnesium (Fig. 2C) for further analyses.

Kinetic Characterization, Product Inhibition, and Lipid Dependence. Cellulose synthase binds the substrate UDP-Glc via its cytosolic glycosyltransferase domain and transfers the activated glucose to the C4 hydroxyl of the nascent cellulose chain, thereby extending the cellulose polymer and forming UDP as a second reaction product (8, 14, 20). Monitoring PttCesA8’s catalytic activity un- + der increasing UDP-Glc concentrations and at a constant Mn2 Fig. 3. Glycosyl linkage analysis of PttCesA8-synthesized cellulose. Gas concentration suggests monophasic Michaelis–Menten kinetics chromatogram corresponding to the separated permethylated alditol ace- with an apparent Km of 30 μM (Fig. 4A). If we assume that k−1 tates. The identity of the derivatives corresponding to 4-linked and terminal is negligible compared with k2, Km can be assimilated to an affinity glucose was confirmed by electron-impact mass spectrometry (Fig. S2). All constant. This affinity compares to an apparent Km for UDP-Glc other peaks of the chromatogram, except for the peak marked with an as- μ R. sphaeroides terisk, correspond to unidentified derivatives that are not alditol acetates, as of about 500 Mfor BcsA (14), perhaps reflecting evidenced by the corresponding fragmentation data. The peak marked with different cytosolic UDP-Glc levels in plant and bacteria. Per- an asterisk corresponds to 4-linked xylose, a known instrument contamina- forming cellulose synthesis reactions at a constant UDP-Glc but tion at the time of analysis. increasing UDP concentrations reveals a profound inhibitory effect of UDP, with an apparent inhibitory constant (Ki)of27μM, matching the enzyme’s affinity (Km) for the substrate UDP-Glc (Fig. cellulose synthase (14). As shown in Fig. 2 A and B, PttCesA8 4B). Other nucleoside diphosphates, such as ADP and GDP, do not continued to form a water-insoluble glucan over 90 min at an opti- inhibit PttCesA8, suggesting that UDP, a reaction product of mal pH and temperature of 7.0 and 35 °C, respectively. The catalytic PttCesA8, rebinds to the active site and inhibits the enzyme com- activity stalls after 90 min of incubation, likely because of product petitively. Indeed, UDP inhibition is overcome at a UDP-Glc con- inhibition or cosedimentation of the synthase and polymeric product. centration about 30 times above the Km, as expected for competitive Processive and nonprocessive inverting glycosyltransferases re- inhibition (Fig. 4C). A similar product inhibition has been observed quire a divalent cation, often a magnesium or manganese ion, for for bacterial cellulose and hyaluronan synthases and is in accordance catalytic activity (3). Accordingly, PttCesA8’s catalytic activity + with a conserved reaction mechanism of these processive family-2 depends on the presence of Mn2 ; no significant enzymatic activity + + glycosyltransferases (14, 21). was observed in the presence of Ca2 or Mg2 , and only modest + activity was seen in the presence of Zn2 (Fig. 2C). To confirm that the observed in vitro catalytic activity is indeed caused by recombinant PttCesA8 and not by a minor contamina- tion with a Pichia protein, we generated a catalytically inactive PttCesA8 mutant. Cellulose biosynthesis requires a general base to deprotonate the acceptor’s hydroxyl group during glycosyl transfer (3, 7, 19). This base is formed by the Asp residue of an invariant TED motif, which we replaced in PttCesA8withanAsn(D676N) to render the enzyme catalytically inactive. The PttCesA8 mutant (PttCesA8*) showed no significant cellulose biosynthetic activity, comparable to material obtained from a mock purification in which Pichia had been transformed with an empty pPICZA vector (Fig. 2C), confirming that the observed biosynthetic activity results from PttCesA8. Further, to establish that the in vitro-synthesized material indeed represents a β-1,4 glucan, the reaction product was hydrolyzed with glucanases specifically degrading β-1,3–, β-1,4–,or α-1,4–linked glucans. Consistent with the formation of authentic cellulose, the PttCesA8-produced material is degraded only by a β-1,4–specific glucanase, a cellulase (Fig. 2D). ’ In addition to testing the polymer s sensitivity to enzymatic Fig. 4. Kinetic analyses of PttCesA8. (A) Titration of UDP-Glc and quantifi- degradation, we also characterized it by permethylation glycosyl cation of the synthesized cellulose. The data were fit to monophasic

linkage analysis. The product obtained from a 60-min synthesis Michaelis–Menten kinetics, yielding a Km of 30 μM. (B) Cellulose biosynthesis reaction contained primarily 1,4-linked glucose together with a in the presence of 30 μM UDP-Glc and increasing concentrations of UDP small amount of terminal glucose (Fig. 3 and Fig. S2). The identity (circles), ADP (triangles), and GDP (squares). UDP inhibits cellulose synthesis μ of the derivatives was confirmed by electron-impact mass spec- with a Ki of about 27 M. (C) UDP competitively inhibits PttCesA8. The trometry (Fig. S2). However, the peak corresponding to terminal catalytic activity of PttCesA8 in the presence of 27 μM UDP and increasing glucose also contained another unidentified compound that was not UDP-Glc concentrations is shown relative to its activity in the absence of a sugar derivative. This compound prevents the calculation of an UDP. (Inset) Lineweaver Burk plot of data shown in A and C.(D) Detergent inhibition of PttCesA8’s catalytic activity. Comparison of PttCesA8’s catalytic accurate degree of polymerization, because the fraction of the peak activity in intact (light gray bars) and solubilized (dark gray bars) proteoli- arising from terminal glucose cannot be determined accurately. posomes formed from S. cerevisiae or E. coli total lipid extracts. (Inset) Western Some PttCesA8 preparations also showed minor product accu- blot of S. cerevisiae (Sc) and E. coli (Ec) proteoliposomes. All experiments were 2+ 2+ mulation in the presence of Mg in addition to the Mn -stimulated performed at least in triplicate; error bars represent SDs from the means.

11362 | www.pnas.org/cgi/doi/10.1073/pnas.1606210113 Purushotham et al. Downloaded by guest on September 27, 2021 affect CesAs catalytic activity. Further, permethylation linkage analysis of cellulose produced by PttCesA8N60 shows a ratio of terminal to internal glucose similar to that observed for the wild- type product (Fig. 3), indicating that the two enzymes produce cellulose polymers of similar lengths (Fig. S5). Although in both analyses the exact amount of terminal glucose is obscured by the presence of either unidentified or 1,3-glucan contaminations, the results suggest a degree of polymerization of at least 25–35 and most likely significantly higher (Fig. 3 and Fig. S5). Fig. 5. PttCesA8’s N terminus is dispensable for catalytic activity. (A)TM topology diagram of PttCesA8 as predicted by TOPCONS (41). The N termini PttCesA8-Synthesized Cellulose Is Partially Resistant to Acid Hydrolysis. of the generated PttCesA8 truncations are indicated. GT, glycosyltransferase. Because of its organization into cellulose micro- and macrofibrils, (B) Comparison of the catalytic activities of all PttCesA8 variants in the ∅ native plant cellulose exhibits a remarkable resistance to acid hydro- presence of 20 mM of the indicated divalent cations or EDTA. denotes lysis. In particular, treatment of cellulosic tissues with Updegraff re- reactions in the absence of additional cations or EDTA. (Inset) Western blot analysis of proteoliposomes containing the indicated PttCesA8 variants. agent [80% acetic acid and concentrated nitric acid at a 10:1 (vol:vol) ratio] (25) has been shown to hydrolyze noncellulosic cell wall materials efficiently as well as isolated glucan chains not organized The heterologously expressed PttCesA8 was solubilized from into fibrils (16, 26). Hence, resistance to acid hydrolysis correlates Pichia membranes and purified in the detergent DDM. Several with a higher-order organization of cellulose. attempts failed to enrich the enzyme in a detergent-solubilized We compared the resistance to acid hydrolysis of PttCesA8 state by the product entrapment method used previously to and bacterial BcsA-synthesized cellulose, the latter forming isolate for example chitin synthases (22). The method relies on noncrystalline cellulose expected to be susceptible to acid hydro- the catalytic activity of the synthase in a detergent-solubilized lysis. To this end, cellulose synthesized in vitro by PttCesA8 and state, so that it cosediments with the synthesized water-insoluble BcsA was resuspended in Updegraff reagent and incubated for polysaccharide. To test whether PttCesA8 is catalytically active in 20 min at 40 °C before quantification. Although the reagent a detergent micelle, we solubilized PttCesA8-containing pro- completely hydrolyzed bacterial cellulose, ∼25% of the PttCesA8- teoliposomes with DDM and analyzed the enzyme’s catalytic synthesized cellulose was resistant to hydrolysis, suggesting a different activity according to our established protocol. As shown in Fig. organization, at least in part, for BcsA- and PttCesA8-produced 4D, PttCesA8 almost completely loses its catalytic activity upon cellulose (Fig. 6A). To confirm that the PttCesA8-produced mate- detergent solubilization of the vesicles, suggesting that enzymatic rial resistant to acid hydrolysis indeed represents cellulose, the activity requires an intact lipid bilayer. This loss of catalytic activity remaining material was washed several times in deionized water is in contrast to bacterial BcsA, which is catalytically active in and subsequently incubated with β-1,3– and β-1,4–specific glucanases several nondenaturing detergents (14). Lack of catalytic activity in of which only the cellulase was able to degrade the product (Fig. 6B). detergents could be caused either by a particular sensitivity of the Interestingly, cellulose produced by PttCesA8 lacking either enzyme toward solubilizing detergents and/or by the disruption of the N-terminal RING-finger (CesA8N60) or the entire N-terminal quaternary structures required for proper function. Despite this cytosolic domain (CesA8N168) was completely hydrolyzed by the dependence, the enzyme exhibits comparable activity in proteoli- Updegraff reagent, perhaps because of the formation of primarily posomes formed from either S. cerevisiae or E. coli total lipid ex- noncrystalline cellulose (Fig. 6A). tracts (Fig. 4D), indicating that no eukaryote-specific lipid, unless copurifying with the enzyme, is required for catalytic activity. PttCesA8 Forms Fiber-Like Cellulosic Structures. The partial resistance of PttCesA8-synthesized cellulose to acid hydrolysis (Fig. 6A), not PttCesA8’s N-Terminal Cytosolic Domain Is Dispensable for Catalytic observed for noncrystalline bacterial cellulose, is consistent with a

Activity. Primarily plant CesAs contain extended cytosolic N higher-order organization of the glucan chains. To analyze the PLANT BIOLOGY termini of about 170 residues that precede the first TM helix organization of the PttCesA8 cellulose product, we visualized the (Fig. 5A). The first ∼65 residues share significant homology with polymers by transmission electron microscopy (TEM) after nega- RING-finger domains, which are specialized Zn-binding domains tive staining. Cellulose was synthesized from proteoliposome- + coordinating two Zn2 ions via a Cys-rich motif. The following ∼100 reconstituted PttCesA8, and the vesicles were solubilized in the residues are less well conserved and have no known function (Fig. S4). detergent Triton X-100, after which the sample was diluted at least Because RING-fingers are often implicated in mediating protein– 100-fold before preparing EM grids. Proteoliposomes containing protein interactions, in particular dimerization (23), and isolated CesA RING-fingers have been shown to form homo- and heterodimers in vitro (24), it is likely that these domains stabilize membrane- embedded CesA oligomers. To test whether PttCesA8’s N terminus is required for catalytic activity, we generated two N-terminally truncated mutants, one devoid of only the RING-finger (CesA8N60, starting with Glu60) and the other missing the entire N-terminal cytosolic domain (CesA8N168, starting with Asn168) (Fig. 5A and Fig. S4). Both mutants exhibit significant catalytic activity relative to the wild-type enzyme, albeit reduced by about 25 and 50% for CesA8N60 and CesA8N168, respectively (Fig. 5B). The lower activity of CesA8N168 likely results from improper folding and/or membrane integra- Fig. 6. PttCesA8-synthesized cellulose is partially acid resistant. (A) Quan- tion, because this mutant expresses at a reduced level and is tification of cellulose resistant to acid hydrolysis. The amount of cellulose resistant to hydrolysis by 80% acetic acid and concentrated nitric acid [10:1 prone to proteolytic degradation. No significant change in catalytic (vol:vol)] is shown relative to the initial cellulosic material produced by each activity was observed for any of the enzymes in the presence of cellulose synthase variant (dark and light gray bars, respectively). (B) Enzy- 2+ 2+ 2+ both Mn and Zn , as compared with Mn only, suggesting matic degradation of the acid-resistant cellulose produced by PttCesA8. GH, 2+ that an intact and Zn -coordinating RING-finger does not directly glycoside hydrolase.

Purushotham et al. PNAS | October 4, 2016 | vol. 113 | no. 40 | 11363 Downloaded by guest on September 27, 2021 (Fig. 3 and Fig. S5). Given also that the mutant’s cellulosic product is particularly sensitive to acid hydrolysis (Fig. 6A), our data suggest that N-terminally truncated PttCesA8 cannot form cellulose microfibrils. Of note, our cellulose biosynthesis assay reports only on the formation of water-insoluble glucans, irrespective of their supramolecular organization. Negative-stain EM, as used in this work, is able to visualize only fibrous material and not noncrystalline cellulose, as confirmed by the observation that BcsA-produced noncrystalline cellulose is not detected (Fig. 7). Taken together, our data suggest that cellulose microfibril formation is not an intrinsic property of CesA and more likely is a secondary effect perhaps arising from a particular CesA quaternary structure supported by the enzyme’s N-terminal cytosolic domain. Fig. 7. PttCesA8 produces cellulose microfibrils in vitro. Cellulose synthesized Discussion by reconstituted PttCesA8 was visualized by negative-stain EM. Shown are representative images of cellulosic material formed by the indicated PttCesA8 CesA functions as polymer synthase and translocase because the constructs. Cellulose microfibrils are seen and in some cases are associated with enzyme synthesizes glucan chains several hundred to thousand globular particles that may be cellulose synthase complexes. Cellulose micro- sugar units long and translocates the polymer across the plasma fibrils originating from a membrane sheet are indicated by a white arrow. membrane during its synthesis. In addition, cellulose polymers Negative controls in the absence of UDP-Glc or with the inactive PttCesA8* synthesized by vascular plants and many other eukaryotic species mutant result in no fiber formation. Infrequent fiber formation is seen with the are assembled into fibrillar structures, thereby establishing their N-terminal truncation mutants. Treatment of the sample with cellulase for physical properties as the load-bearing component of the cell wall. 10 min before grid preparation results in the loss of cellulose fibers. BcsA- produced cellulose was not detectable. Plant CesAs have been shown to interact with several components, including cytosolic microtubule-binding proteins, and also with membrane-integrated or -attached factors, such as the wild-type PttCesA8 produced fiber-like polymers several microme- β-1,4 glucanase KORRIGAN (33). However, the cellulose bio- ters long only when incubated in the presence of substrate. Control synthetic activity of recombinantly expressed and reconstituted reactions in the absence of UDP-Glc did not contain any polymeric CesA8 from hybrid aspen demonstrates that no other plant-specific material (Fig. 7). Some samples contained apparently incompletely proteinaceous or lipidic factors are required for its catalytic activity. solubilized membrane fragments, clearly visible by negative-stain Recent advances in bacterial cellulose biosynthesis provide EM, from which cellulose fibrils originated, suggesting that mi- detailed insights into how the synthase elongates the nascent crofibril formation occurs directly after the release of the nascent cellulose polymer one glucose unit at a time with translocation glucans from their site of synthesis. steps between each round of polymer extension (8). However, it The apparent cellulose microfibrils were produced only by is currently unknown how cellulose biosynthesis is initiated and wild-type PttCesA8 and not by the catalytically inactive PttCesA8* in particular whether a specific primer is required to form the mutant, confirming that fiber formation indeed is a result of first acceptor of glycosyl transfer. Several proposed models for PttCesA8’s catalytic activity (Fig. 7). Additionally, the synthesized fibers bacterial and plant cellulose biosynthesis include the in- could be degraded with cellulase, resulting in almost complete loss volvement of lipid-linked oligosaccharides or glycolipids, such as of visible fibers within 10 min of incubation (Fig. 7 and Fig. S6). sitosterol-glucoside, which has been found only in plants so far (34–36). However, the catalytic activity of Pichia-expressed Analysis of frozen hydrated fibers by cryo-electron microscopy Ptt (cryoEM) suggests a diameter of 48 ± 10Å (Fig. S7), which is in CesA8 suggests that the enzyme either initiates cellulose biosynthesis in vivo using an expression host-provided factor [as agreement with diameter estimates for cellulose microfibrils observed for Rhodobacter BcsA upon expression in E. coli (14)] or obtained from spruce wood and celery collenchyma (27, 28). Assuming that cellulose biosynthesis initiates in vitro upon incubation with a stacking distance of about 5 Å between glucan chains within a + UDP-Glc and Mn2 . Based on the Rhodobacter BcsA structure, cellulose microfibril, the estimated width of the PttCesA8-produced – priming of cellulose biosynthesis with monomeric glucose (derived fiber is sufficient to accommodate at least 18 24 glucan chains. Indeed, from UDP-Glc hydrolysis) seems likely, as recently discussed (6, 37). comparable cellulose fibers have been synthesized from solubilized Ptt Physcomitrella patens In contrast to BcsA, CesA8-synthesized cellulose appears as CesA5 (17), with a similar estimated diameter of long fibers that are partially resistant to acid hydrolysis. It is 20–30 Å, as well as from blackberry membrane extracts (16). ∼ tempting to speculate that in vivo these microfibrils could co- Some fibers are capped at one end by globular structures 250 Å alesce to form macrofibrils, similar to those isolated from native in diameter (Fig. 7). Similar structures have been observed on tissues and giving rise to X-ray powder diffraction (28). The lim- cellulose fibers produced by CesAs solubilized from blackberry ited amount and random orientation of the PttCesA8-produced P patens and . membranes and most likely represent polymer- microfibrils currently preclude a similar analysis of our product. attached CesAs (16, 17). The strong interaction of cellulose synthase However, the observations that recombinant PttCesA8 forms Rhodobacter with the nascent cellulose polymer is exemplified by cellulose microfibrils and BcsA does not suggest that microfibril E coli BcsA, which, upon heterologous expression in . and detergent formation is caused by CesA-specific features. An attractive solubilization, copurifies with a cellulose chain (7). The width of the model is that CesA oligomerization suffices to drive glucan chain fibril-attached particle is consistent with size estimates for plant alignment, consistent with the large globular densities capping CesA oligomers, referred to as “rosettes,” observed by freeze-frac- some of the cellulose microfibrils (Fig. 7). The observations that ture EM (11, 29, 30). CesA rosettes have recently been interpreted N-terminally truncated PttCesA8s are catalytically active but fail as hexamers of CesA trimers, yielding a total of 18 CesA molecules to form cellulose microfibrils further supports this hypothesis. + per complex and, accordingly, a maximum of 18 glucan chains per CesA’s N-terminal Zn2 -binding region has strong similarity to cellulose microfibril (31, 32). RING-fingers and likely mediates CesA–CesA interactions. Interestingly, cellulose produced by the N-terminally truncated Further, plant CesAs contain two prominent insertions within PttCesA8s contained very little to no fibrous material (Fig. 7), de- the evolutionarily conserved glycosyltransferase domain, referred spite significant in vitro catalytic activity (Fig. 5B) and an apparent to as “plant-conserved” and “class-specific” regions (38), which have polymer length similar to that obtained from wild-type PttCesA8 been proposed to be involved in CesA oligomerization as well (9).

11364 | www.pnas.org/cgi/doi/10.1073/pnas.1606210113 Purushotham et al. Downloaded by guest on September 27, 2021 CesA oligomers (rosettes) contain different isoforms; for example, Methods in Arabidopsis and Populus CesA1,3,and6areinvolvedinprimary Populus tremula × tremuloides CesA8 was expressed as a C-terminally His-tagged and CesA4, 7, and 8 are involved in secondary cell wall biosynthesis species in P. pastoris and was purified in the detergent LysoFoscholine-Ether14 by (1). Stoichiometry estimates of CesA rosettes formed during primary metal-affinity chromatography. The purified protein was reconstituted into proteoliposomes, in which it synthesized cellulose upon incubation with the substrate or secondary cell wall formation suggest an equal number of isoforms + UDP-glucose and Mn2 cations. The synthesized polymer was quantified by scin- per complex (39, 40). Although the arrangements and functions 3 of the individual CesAs in a rosette are currently unknown, our tillation counting upon incorporation of trace amounts of H-labeled glucose as described (14). The synthesis of authentic cellulose was confirmed by en- data demonstrate that a single isoform suffices to form a cellulose zymatic degradation and permethylation glycosyl linkage analysis as described microfibril. This finding raises the possibility that CesA rosettes for bacterial cellulose synthase (14). Uranyl formate-stained electron micro- represent higher-order complexes of catalytically active CesA graphs were obtained from in vitro-synthesized cellulose after the proteoli- homooligomers, perhaps preformed dimers or trimers, as recently posomes were solubilized in the detergent Triton X-100. Detailed methods are observed for an isolated glycosyltransferase domain from Arabidopsis provided in SI Methods. CesA1 (31). Rosette formation then could be driven by interactions between preformed CesA homooligomers of different isoforms. ACKNOWLEDGMENTS. We thank Chao Fang and Caitlin Hubbard for assis- The reconstitution of cellulose biosynthesis from heterolo- tance with negative-stain EM and total internal reflection fluorescence Ptt microscopy, respectively. Protein MS-MS fingerprinting was performed gously expressed and purified CesA8 provides a powerful tool by the Taplin Mass Spectrometry Facility, Harvard Medical School. P.P., to delineate many crucial steps during cellulose microfibril for- S.H.C., M.K., B.T.N., and J.Z. are supported by the Center for Lignocellulose mation. These steps include interaction studies of PttCesA8 and Structure and Formation, Energy Frontier Research Center, US Department its cellulosic product with other proteins and complex carbohy- of Energy, Office of Science Award DE-SC0001090. S.M.D.-M. and V.B. are supported by the Australian Research Council Centre of Excellence drates, thereby providing an in vitro platform for investigating in Plant Cell Walls and matching funding from the Royal Institute of the many steps implicated in plant cell wall formation. Technology, Stockholm.

1. McFarlane HE, Döring A, Persson S (2014) The cell biology of cellulose synthesis. Annu 21. Tlapak-Simmons VL, Baron CA, Weigel PH (2004) Characterization of the purified Rev Plant Biol 65:69–94. hyaluronan synthase from Streptococcus equisimilis. Biochemistry 43(28):9234–9242. 2. Bi Y, Hubbard C, Purushotham P, Zimmer J (2015) Insights into the structure and 22. Kang MS, et al. (1984) Isolation of chitin synthetase from Saccharomyces cerevisiae. function of membrane-integrated processive glycosyltransferases. Curr Opin Struct Purification of an enzyme by entrapment in the reaction product. J Biol Chem Biol 34:78–86. 259(23):14966–14972. 3. Lairson LL, Henrissat B, Davies GJ, Withers SG (2008) Glycosyltransferases: Structures, 23. Liew CW, Sun H, Hunter T, Day CL (2010) RING domain dimerization is essential for functions, and mechanisms. Annu Rev Biochem 77:521–555. RNF4 function. Biochem J 431(1):23–29. 4. Newman RH, Hill SJ, Harris PJ (2013) Wide-angle x-ray scattering and solid-state nu- 24. Kurek I, Kawagoe Y, Jacob-Wilk D, Doblin M, Delmer D (2002) Dimerization of cotton clear magnetic resonance data combined to test models for cellulose microfibrils in fiber cellulose synthase catalytic subunits occurs via oxidation of the zinc-binding mung bean cell walls. Plant Physiol 163(4):1558–1567. domains. Proc Natl Acad Sci USA 99(17):11109–11114. 5. Jarvis MC (2013) Cellulose biosynthesis: Counting the chains. Plant Physiol 163(4): 25. Maji S, Mehrotra R, Mehrotra S (2013) Extraction of high quality cellulose from the 1485–1486. stem of Calotropis procera. South Asian J Exp Biol 3(3):113–118. 6. McNamara J, Morgan JLW, Zimmer J (2015) A molecular description of cellulose 26. Updegraff DM (1969) Semimicro determination of cellulose in biological materials. biosynthesis. Annu Rev Biochem 84:17.11–17.27. Anal Biochem 32(3):420–424. 7. Morgan JL, Strumillo J, Zimmer J (2013) Crystallographic snapshot of cellulose syn- 27. Thomas LH, et al. (2013) Structure of cellulose microfibrils in primary cell walls from thesis and membrane translocation. Nature 493(7431):181–186. collenchyma. Plant Physiol 161(1):465–476. 8. Morgan JL, et al. (2016) Observing cellulose biosynthesis and membrane translocation 28. Fernandes AN, et al. (2011) Nanostructure of cellulose microfibrils in spruce wood. in crystallo. Nature 531(7594):329–334. Proc Natl Acad Sci USA 108(47):E1195–E1203. 9. Sethaphong L, et al. (2013) Tertiary model of a plant cellulose synthase. Proc Natl 29. Herth W (1983) Arrays of plasma-membrane “rosettes” involved in cellulose micro- Acad Sci USA 110(18):7512–7517. fibril formation of Spirogyra. Planta 159(4):347–356. 10. Slabaugh E, Davis JK, Haigler CH, Yingling YG, Zimmer J (2014) Cellulose synthases: 30. Roberts AW, Roberts EM, Haigler CH (2012) Moss cell walls: Structure and bio- New insights from crystallography and modeling. Trends Plant Sci 19(2):99–106. synthesis. Front Plant Sci 3:166. 11. Kimura S, et al. (1999) Immunogold labeling of rosette terminal cellulose-synthesizing 31. Vandavasi VG, et al. (2016) A structural study of CESA1 catalytic domain of Arabidopsis complexes in the vascular plant Vigna angularis. Plant Cell 11(11):2075–2086. cellulose synthesis complex: Evidence for CESA trimers. Plant Physiol 170(1):123–135. 12. Brown RM (2003) Cellulose structure and biosynthesis: What is in store for the 21st 32. Nixon BT, et al. (2016) Comparative structural and computational analysis supports –

century? J Polym Sci A Polym Chem 42(3):487 495. eighteen cellulose synthases in the plant cellulose synthesis complex. Sci Rep 6:28696. PLANT BIOLOGY 13. Giddings TH, Jr, Brower DL, Staehelin LA (1980) Visualization of particle complexes in 33. Vain T, et al. (2014) The cellulase KORRIGAN is part of the cellulose synthase complex. the plasma membrane of Micrasterias denticulata associated with the formation of Plant Physiol 165(4):1521–1532. cellulose fibrils in primary and secondary cell walls. J Cell Biol 84(2):327–339. 34. Matthysse AG, Thomas DL, White AR (1995) Mechanism of cellulose synthesis in 14. Omadjela O, et al. (2013) BcsA and BcsB form the catalytically active core of bacterial Agrobacterium tumefaciens. J Bacteriol 177(4):1076–1081. cellulose synthase sufficient for in vitro cellulose synthesis. Proc Natl Acad Sci USA 35. Peng L, Kawagoe Y, Hogan P, Delmer D (2002) Sitosterol-beta-glucoside as primer for 110(44):17856–17861. cellulose synthesis in plants. Science 295(5552):147–150. 15. Cifuentes C, Bulone V, Emons AMC (2010) Biosynthesis of callose and cellulose by 36. Grillitsch K, et al. (2014) Isolation and characterization of the plasma membrane from detergent extracts of tobacco cell membranes and quantification of the polymers the yeast Pichia pastoris. Biochim Biophys Acta 1838(7):1889–1897. synthesized in vitro. J Integr Plant Biol 52(2):221–233. 37. Morgan JLW, McNamara JT, Zimmer J (2014) Mechanism of activation of bacterial 16. Lai-Kee-Him J, et al. (2002) In vitro versus in vivo cellulose microfibrils from plant cellulose synthase by cyclic di-GMP. Nat Struct Mol Biol 21(5):489–496. primary wall synthases: Structural differences. J Biol Chem 277(40):36931–36939. 38. Vergara CE, Carpita NC (2001) Beta-D-glycan synthases and the CesA gene family: 17. Cho SH, et al. (2015) In vitro synthesis of cellulose microfibrils by a membrane protein Lessons to be learned from the mixed-linkage (1–>3),(1–>4)beta-D-glucan synthase. from protoplasts of the non-vascular plant Physcomitrella patens. Biochem J 470(2): Plant Mol Biol 47(1-2):145–160. 195–205. 39. Hill JL, Jr, Hammudi MB, Tien M (2014) The Arabidopsis cellulose synthase complex: A 18. Gray MC, Converse AO, Wyman CE (2003) Sugar monomer and oligomer solubility: proposed hexamer of CESA trimers in an equimolar stoichiometry. Plant Cell 26(12): Data and predictions for application to biomass hydrolysis. Appl Biochem Biotechnol 4834–4842. 105 -108:179–193. 40. Gonneau M, Desprez T, Guillot A, Vernhettes S, Höfte H (2014) Catalytic subunit 19. Yang H, Zimmer J, Yingling YG, Kubicki JD (2015) How cellulose elongates–a QM/MM stoichiometry within the cellulose synthase complex. Plant Physiol 166(4):1709–1712. study of the molecular mechanism of cellulose polymerization in bacterial CESA. 41. Bernsel A, Viklund H, Hennerdal A, Elofsson A (2009) TOPCONS: Consensus prediction J Phys Chem B 119(22):6525–6535. of membrane protein topology. Nucleic Acids Res 37(Web Server issue):W465–468. 20. Brown C, Leijon F, Bulone V (2012) Radiometric and spectrophotometric in vitro assays 42. Djerbi SAH, et al. (2004) Identification and expression analysis of genes encoding of glycosyltransferases involved in plant cell wall carbohydrate biosynthesis. Nat putative cellulose synthases (CesA) in the hybrid aspen, Populus tremula (L.) x Protoc 7(9):1634–1650. P-tremuloides (Michx.). Cellulose 11:301–312.

Purushotham et al. PNAS | October 4, 2016 | vol. 113 | no. 40 | 11365 Downloaded by guest on September 27, 2021