DOI: 10.1126/science.1227491, 1055 (2012);338 Science et al.Shi-You Ding

Enzymatic Digestibility?How Does Plant Cell Wall Nanoscale Architecture Correlate with

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15. Materials and methods are available as supplementarymaterials on Science Online.

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SC-RR-710714, Sandia Nat. Labs (1972).18. H. J. Melosh, Meteorit. Planet. Sci. 42, 2079 (2007).19. S. Ida, R. M. Canup, G. R. Stewart, Nature 389, 353 (1997).20. E. Kokubo, J. Makino, S. Ida, Icarus 148, 419 (2000).21. M. M. M. Meier, A. Reufer, W. Benz, R. Wieler,

Annual Meeting of the Meteoritical Society LXXIV,abstr. 5039 (2011).

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Acknowledgments: SPH simulation data are contained intables S2 to S5 of the supplementary materials. Financialsupport for this project was provided by the NASA LunarScience Institute and by NASA’s LASER program.

Supplementary Materialswww.sciencemag.org/cgi/content/full/science.1226073/DC1Materials and MethodsFigs. S1 and S2Tables S1 to S5References (26–36)

14 June 2012; accepted 1 October 2012Published online 17 October 2012;10.1126/science.1226073

How Does Plant Cell WallNanoscale Architecture Correlatewith Enzymatic Digestibility?Shi-You Ding,1*† Yu-San Liu,1* Yining Zeng,1 Michael E. Himmel,1

John O. Baker,1 Edward A. Bayer2

Greater understanding of the mechanisms contributing to chemical and enzymatic solubilization ofplant cell walls is critical for enabling cost-effective industrial conversion of cellulosic biomass tobiofuels. Here, we report the use of correlative imaging in real time to assess the impact ofpretreatment, as well as the resulting nanometer-scale changes in cell wall structure, upon subsequentdigestion by two commercially relevant cellulase systems. We demonstrate that the small, noncomplexedfungal cellulases deconstruct cell walls using mechanisms that differ considerably from those of thelarger, multienzyme complexes (cellulosomes). Furthermore, high-resolution measurement ofthe microfibrillar architecture of cell walls suggests that digestion is primarily facilitated byenabling enzyme access to the hydrophobic cellulose face. The data support the conclusion that idealpretreatments should maximize lignin removal and minimize polysaccharide modification,thereby retaining the essentially native microfibrillar structure.

Modern biorefineries are being devel-oped to produce transportation fuelsfrom plant biomass using sustainable

technologies that also benefit the environmentby reducing greenhouse gas emissions (1). Themajor challenges facing this industry are thehigh cost of pretreatment and the low efficiencyof enzymatic hydrolysis of plant cell wall poly-saccharides to sugars. Further improvement ofthese processes is contingent on deeper under-standing of biomass structure and chemistry, aswell as improved understanding of the molecu-lar mechanisms of biomass deconstruction (2).

Despite renewed interest in biomass deg-radation, there is little agreement about whichplant cell wall features most affect digestibility bymicrobes and cellulolytic enzymes. The activitiesof cellulolytic enzymes are usually characterizedby assay against purified or highly modified crys-talline or amorphous celluloses, as well as solu-ble substrates (3). Actual plant cell walls, however,are complex nanocomposites containing networksof cellulose fibrils and complex “matrixing” poly-

mers. The overall performance of biomass sac-charification may be attributed to the synergisticaction of many complementary enzymes—includinga variety of cellulases, hemicellulases, and ac-cessory enzymes (4)—which makes it difficult tostudy one factor at a time. Traditional solutionmethods have suffered from the classical ensem-ble average limitation presented by analysis ofthese mixtures of complex biomass; therefore, thedata gathered are sometimes inconclusive and,in part, contradictory. To overcome these prob-lems, we visualized the action of these enzymesystems on untreated and delignified plant cellwalls under controlled digestion conditions inreal time with the use of a multimodal micros-copy suite, including bright-field light microsco-py, confocal laser scanning microscopy (CLSM),two-color stimulated Raman scattering (SRS) mi-croscopy, and atomic force microscopy (AFM).The microscopes were custom-constructed to al-low correlative imaging of the same biomass sam-ple under near-physiological conditions and athigh chemical (5, 6) and spatial (7, 8) resolu-tions at the tissue, cellular, and molecular levels.We examined two naturally existing enzyme sys-tems of commercial relevance for saccharificationof lignocellulosic biomass: (i) the secretome of theanaerobeClostridium thermocellum, which is rep-resentative of multienzyme bacterial cellulosomes(9), and (ii) a commercially available blended en-zymemixture (CellicCTec2,Novozymes,Bagsværd,

Denmark) derived from the fungus Trichodermareesei (Hypocrea jecorina), which is represent-ative of the fungal or “free” cellulases (10). Weused green fluorescent protein (GFP)–taggedcarbohydrate-binding modules (CBMs) to iden-tify exposed cellulose surfaces and green dye(Alexa Fluor 488, Invitrogen, Carlsbad, CA)–labeled enzymes to examine overall cell wall ac-cessibility to cellulases. TrCBM1 derived fromT. reesei cellobiohydrolase I (CBH I or Cel7A)(11) and CtCBM3 derived fromC. thermocellumcellulosomal scaffoldin protein (CipA) (12) spe-cifically recognize the planar face of crystallinecellulose (8, 13, 14) and play a critical role in thehydrolysis of crystalline cellulose (15, 16).

Naturally senescent, dried corn (Zea mays L.)stover stem internode sections served as an ex-ample of plant cell wall material. The transverseview of the untreated maize stem represents thetypical tissue structure of monocotyledons withscattered vascular bundles (VBs) surroundedby parenchyma cells (fig. S1, A and B). In thiscontext, cell walls in mature plants are generallyclassified as one of three types: (i) The pri-mary walls (PWs) are thin (~100 nm) and ex-panded, but are neither elongated nor lignified.(ii) Parenchyma-type secondary walls (pSWs)are found in large parenchyma tissue betweenthe VBs, which are elongated or expanded andpartially lignified in the secondarily thickenedwall (~1 to 2 mm). (iii) Sclerenchyma-type SWs(sSWs) are elongated and fully lignified in theheavily thickened (~5 to 10 mm) SW. The in-nermost side of a sSW is often covered by “warts,”which mainly contain condensed lignin-like poly-phenols (17). Note that the materials used inthis study are from dead plants; cell walls in aliving plant may be more structurally complexand diversified.

Using two-color SRS microscopy, wherethe Raman signal at 1600 cm−1 (aromatic-ringbreathing modes) represents primarily ligninand the 2900 cm−1 (C-H stretch) band repre-sents primarily polysaccharides (18), we foundthat the lignin and polysaccharide contents insSWs were higher than those in pSWs (fig.S1, C to E) and that the polysaccharide con-tent was generally higher in SWs than in PWs.These observations are in agreement with gen-eral plant anatomy.

The presence of lignins, a group of highlybranched phenylpropanoid polymers found interrestrial plants, is generally considered to beone of the most important limiting factors in

1Biosciences Center, National Renewable Energy Laboratory,Golden, CO 80401, USA. 2Department of Biological Chem-istry, The Weizmann Institute of Science, Rehovot 76100,Israel.

*These authors contributed equally to this work.†To whom correspondence should be addressed. E-mail:shi.you.ding@nrel.gov

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the enzymatic cell wall saccharification process(19). Whether lignins affect enzyme digestibilityby physically impeding or nonspecifically absorb-ing enzymes remains open to debate. To addressthis question, we used acid chlorite to producedelignified cell walls. This delignification processhas been widely used to produce holocellulose(20). At low temperature, acid chlorite oxidizesphenolic compounds (mainly lignins) in cell wallswithout altering cellulose and with minimum ef-fect on associated hemicelluloses (5, 21).

As anticipated, we confirmed that accessibil-ity of untreated cell walls to CBMs and enzymebinding exhibits a strong negative correlationwith their lignin content. All CBMs and enzymesbind strongly to nonlignified PWs and moreweakly to pSWs. Binding to the condensed lig-nin “warty layer” in sSWs is negligible, indicat-ing that nonspecific adsorption of processedlignin to enzyme reported previously (22) wasnot observed in the case of a native lignin ex-amined in situ. Lignin removal enhanced overallbinding of all probes to lignified walls (i.e., pSWs

and sSWs) (Fig. 1), following the trend: fungalcellulases (greatest enhancement), cellulosomes,TrCBM1, and then CtCBM3 (least enhancement).Accessibility of pSWs and sSWs to enzymes wasmore strongly enhanced than that to CBMs, whichcould generally be attributed to increased accessi-bility of hemicelluloses to enzymes after removalof blocking lignins. Further imaging of deligni-fied pSWs by CLSM showed that cellulosomesprimarily attached to the surface of the cell walls,the cell corners, and the plasmodesmata (Fig. 1,B and C, and fig. S2A). Fungal cellulases notonly bound at these surfaces, but also penetratedinside the SW from its innermost side (Fig. 1,D and E, and fig. S2B), suggesting that delig-nification may render the cell wall more acces-sible to fungal cellulases.

After delignification, we found that the lig-nin signal at 1600 cm−1 was eliminated and the2900-cm−1 signal was slightly reduced, suggest-ing that hemicelluloses are generally resistant toacid chlorite treatment at room temperature (Fig.2, A to E).

The fungal cellulases and cellulosomes thatwe used in this study contain mixtures of en-zymes specific for degradation of different poly-saccharide components of cell walls. The digestiontemperatures used here were optimal for the fun-gal (38°C) and cellulosomal (55°C) enzymes. Inuntreated cell walls, both enzyme systems readilydigested PWs only (Fig. 2, F, G, J, and K), andSRS microscopy also revealed minor digestionof pSWs (Fig. 2E and fig. S3). Overall digest-ibility was strongly negatively correlated with lignincontent. Complete degradation was not observedin naturally lignified walls (i.e., pSWs and sSWs),even with 100-fold greater concentrations of en-zyme and extended periods of time (7 days).

After delignification, all cell walls were com-pletely digested with the original level of cellu-lase protein loading (Fig. 2, H and L). Despitethe difficulties of quantitatively determining en-zyme digestion rates by microscopy alone, wediscovered that fungal cellulases acted approx-imately five times faster than cellulosomes againsteither untreated or delignified cell walls, based on

Fig. 1. Confocal laserscanning microscopy ofcell walls exposed to GFP-CBMs and dyed cellulases.All probes are shown ingreen. (A) Transverse sec-tion of VB area. CBMsspecifically recognize cel-lulose, which is highlyaccessible in PWs but lessaccessible in pSWs (andis not accessible in sSWs).Autofluorescence (red) andoverlay images highlightthe negative correlationbetween probe bindingand autofluorescence. De-lignification significantlyincreases cell wall accessi-bility to enzymes (pairedt test, *P < 0.05). Histo-grams showing relativefluorescence intensity areexpressed as percentagesof fluorescence comparedwith the intensity of thelabeled PW, which is des-ignated as 100%. Delig-nified pSWs in the rindarea were imaged inhigher magnification. (B)Cellulosomes bound tothe pSW innermost sur-face, the cell corners (CC),and the plasmodesmata(P). (C) Composite-averageprofile of line scans acrossthe SW, as illustrated bythe dotted line in (B).(D) Fungal cellulases pen-etrating into the pSW fromthe innermost surface. (E) Composite-average profile of line scans, as illustrated by the dotted line in (D) showing further penetration (red arrowhead). Dataare mean T SD based on analysis of five images (A, C, and E). Scale bars: 50 mm (A), 5 mm (B and D).

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the percentage loss of SRS signals at 2900 cm−1.The time needed for complete digestion of thedelignified walls apparently depends on wallmass concentration and not on wall types (Fig.2, I and M). Thinner and less dense walls weredigested rapidly, suggesting that the chemicalfeatures of the polysaccharides existing in dif-ferent cell wall types may have less impact onoverall digestibility than expected, at least in thecase of acid chlorite–treated biomass, in whichlignins have been selectively oxidized.

From experiments at room temperature,we made the qualitative observations that thecellulosomes first separated the walls from thecompound middle lamella (CML) and then frag-mented the walls into segments (Fig. 3A andmovie S1), whereas fungal cellulases dissolved theentire wall in a uniform manner (Fig. 3B andmovie S2). During the course of digestion byfungal cellulases, the CML remained mostlyintact, suggesting that digestion occurred fromthe innermost side of the cell walls, thus sup-porting the binding data (Fig. 1D).

We used AFM for real-time imaging of pSWdigestion at the microfibril level. In the untreated

cell wall, only a few surface microfibrils weredegradable by either of the enzyme systems.After delignification, the cellulosomes appearedto peel off individual microfibrils from the cellwall surface (Fig. 3C and movie S3), whereasthe fungal cellulases penetrated inside micro-fibril networks and made digestion holes or pits(Fig. 3D and movie S4). For quantitative anal-ysis, the surface roughness was expressed byrelative Z ranges of AFM images. In the case ofcellulosomes, the roughness remained unchanged(Fig. 3E), whereas digestion by fungal cellulasesincreased the roughness dramatically (Fig. 3F).Individual microfibrils were digested within afew minutes by each of the enzyme systems. Be-cause AFMmovies were acquired within a 1-mm2

scan area, the length of the microfibrils isunknown; therefore, the digestion rate cannot beestimated unambiguously. The wall structureunder buffer was similar to that in air with minordifferences, consisting primarily of straighteningand debundling surface fibrils (fig. S4).

Cellulose represents ~70% of the total cellwall polysaccharides, and its accessibility to en-zymes determines the overall digestibility. We

have previously reported that CBH I binds toand hydrolyzes the hydrophobic planar faces ofpurified cellulose crystals (8). To further iden-tify the cell wall architectural features that maycontribute to differences in accessibility to anddigestibility by cellulases, we used nanometer-scale AFM to image untreated and delignifiedcell walls.

As previously reported, we observed threetypes of fibrillar structures (7, 23): Ribbon-likemacrofibrils, consisting of a number of celluloseelementary fibrils (CEFs), exist in PWs (fig. S5)and on the pSW surface (Fig. 4A and fig. S6).Microfibrils, which contain one CEF and asso-ciated hemicelluloses, are predominant in pSWsand sSWs (fig. S7). Although smaller CEFs,such as the 24-chain diamond-shaped model(24), have been proposed, we favor the 36-chainmodel, primarily based on microfibril size mea-sured directly in the current study and on twoprimary assumptions: (i) that cell wall celluloseis mostly crystalline and (ii) that the CBM bind-ing face recognizes the cellulose planar face in-volving three chains (14, 25). Therefore, packingof cellulose Ib (26) would allow three chains on

Fig. 2. Stimulated Raman scattering (SRS) microscopy of untreated cell wallsshowing (A) polysaccharides at 2900 cm−1 (blue) and (B) lignins at 1600 cm−1

(red). After delignification, (C) the signal at 2900 cm−1 is slightly reduced, and(D) the 1600 cm−1 signal is eliminated. (E) Histograms of SRS signals in un-treated and delignified cell walls and enzyme-digested residues. Bright-fieldlight images of untreated cell walls before (F) and after (G) digestion bycellulosomes for 15 hours at 55°C showing that only the PWs are completelydigested. (H) Delignified walls digested by the cellulosomes for 17 hours at55°C, showing that all cell walls are digested. (I) Relative digestion rates by

cellulosomes. (J and K) Untreated cell walls before (J) and after (K) digestionby fungal cellulases for 3 hours at 38°C, showing that only the PWs aredigested. (L) Delignified cell walls are completely digested by fungal cellu-lases in 8 hours at 38°C. (M) Relative digestion rates by fungal cellulases.Color bars in (A) to (D) show the lock-in amplifier signals of SRS. Signals thatare not detectable are indicated by asterisks in boxes in (E), (I), and (M).Relative digestion rates were calculated based on the percentage loss of SRSsignal at 2900 cm−1 per hour (I and M). Data are mean T SD based on triplicatemeasurements (E, I, and M). Scale bar: 50 mm (A to L).

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the planar faces (100 and –100), supporting the36-chain model.

High-resolution AFM images also allowedprecise measurement of the size of fibrillar struc-tures. Isolated microfibrils (Fig. 4A) that ap-peared twisted were found occasionally on thedry cell wall surface. Note that this twisting ef-fect was not found under aqueous conditions,suggesting that it may be caused by dehydra-tion. In such cases, the AFM line-scan profilesof both vertical (Fig. 4B) and horizontal (Fig.4C) orientations could be measured on the sameCEF. Our measurement of a macrofibril contain-ing two CEFs suggested that they are horizon-tally associated through their hydrophilic faces(Fig. 4, D and E). Individual microfibrils in pSWswere visualized as sharp edges with bridges par-tially covering the area between them (Fig. 4, Aand D, and fig. S6E), in which the CEF appearedto be primarily vertically oriented. Macrofibrilsplitting and CEF rotation from horizontal tovertical may occur during cell elongation or ex-pansion (fig. S8).

Quantitative analysis using AFM scan pro-files of macrofibrils, microfibrils, and CEFs (Fig.4F) showed that the widths are slightly larger[3.3 T 0.5 nm (vertical) and 5.7 T 0.9 nm (hori-zontal)] than those predicted from the 36-chainmodel (7) [i.e., 3.2 nm (vertical) by 5.3 nm (hori-zontal)], which may be caused by a combinedeffect of AFM tip broadening and CEF asso-ciation with matrix polymers, especially in sSWs(fig. S7C). The microfibrils in the SW appearedto form parallel layers, observed in transversesections (fig. S7E) and the broken surface insSWs after delignification (fig. S7I). Cell wallporosity (open space) is measured by three dis-tances between fibrils (5 to 20 nm), layers ofmicrofibrils (10 to 20 nm) (7), and wall lamellaegaps (50 to 100 nm) appearing in the SWs (fig.S7B). Delignification did not substantially altermicrofibril structure and arrangement, except forthe reduction of matrix polymers of the sSW(fig. S7, G and H).

Schematic structures of the PW and thesandwich-like SW in pSWs and sSWs containinglayers of microfibrils and lignin-polysaccharidesare illustrated in Fig. 4G. Here, we combine theconcepts suggested by the AFM and digestiondata, which include the bundling of microfibrilsinto macrofibrils in PWs and the natural avail-ability of cellulose planar faces in PWs.

These observations of cell wall architectureand microfibril structure are consistent with ac-cessibility to (Fig. 1) and digestibility by (Fig. 2)enzymes, suggesting that the hydrophobic pla-nar face of cellulose is the preferred binding faceand thus is critical for enzyme access. In PWs,the CEF hydrophobic faces in macrofibrils areexposed to allow accessibility to cellulases, asconfirmed by the strong binding of CBMs andcomplete digestibility by cellulases. In pSWs, thesmall surface macrofibrils and microfibrils areaccessible, but microfibrils inside the wall areblocked by the lignin-polysaccharide complex

between microfibril layers (Fig. 4G), resulting indiminished CBM binding and limited digestibilityby enzymes. In sSWs, lignins form an additionalbarrier on the innermost side, the condensed lig-nin warty layer (17), that blocks accessibility ofmicrofibrils to enzymes (Fig. 1), resulting in nodigestibility of sSWs (Fig. 2). These observationsare also supported by studies on cell wall devel-opment, which have indicated that SW thicken-ing begins while the cell is still elongating (27).In later stages of elongation, SW deposition isaccompanied by a lignification process that maycause a decrease and eventual cessation of cellelongation. pSWs are not lignified until cessationof vegetative growth in grassy plants; conse-quently, these walls are completely digestible ifharvested during vegetative growth (28).

Acid chlorite treatment effectively removeslignins in the SW (Fig. 2 and fig. S7H) and the

warty layer in sSW (fig. S7F), thereby exposingmicrofibrils (fig. S7, G to I) to enzyme access,resulting in near-complete digestion of all cellwalls. After lignin removal in the SWs, acces-sibility of the cellulose planar face is then deter-mined by wall porosity. Under the conditions ofthis study, we observed enhanced digestibilityby fungal cellulases due to their penetration intothe pore structure of microfibril networks (Fig. 3Dand movie S4). In contrast, the larger cellulosomecomplexes could only penetrate the larger walllamella gaps (fig. S7B), resulting in fragmen-tation of walls (Fig. 3A and movie S1). Theadvantageous degradation properties exhibited bythe fungal cellulases on cell walls may be com-promised when digesting purified forms of crys-talline cellulose, such as Avicel PH101 (Sigma,St. Louis, MO) (29), in which the porous archi-tecture of the native cellulose microfibril network

Fig. 3. Delignified pSWs imaged in real time during digestion at room temperature. Bright-field lightmicroscopy of a transverse section digested (A) by cellulosomes for 7 days, showing wall fragmentation(white arrow), and (B) by fungal cellulases for 10 hours, showing wall dissolving. White arrows in (B)indicate the wall’s innermost side. AFM imaging of a single pSW surface digested (C) by cellulosomesfor 13 hours, showing peeling-off of individual microfibrils (white arrows), and (D) by fungal cellulasesfor 2 hours, showing penetration (white arrows). Images in (A) to (D) were taken from movies S1, S2,S3, and S4, respectively. Relative Z ranges of AFM images were recorded as changes of vertical distanceduring digestion by (E) cellulosomes (movie S3), showing conserved roughness, and by (F) fungalcellulases (movie S4), showing increasing roughness. Black arrows indicate the time points of enzymeaddition. Color bars in (C) and (D) represent the scale of the AFM height images. Scale bars: 50 mm(A and B), 200 nm (C and D).

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has been completely destroyed during its prepa-ration process. Similarly, it has been reported thatbiomass digestibility was notably reduced uponnear-complete removal of both lignin and hemi-celluloses (21). In contrast, the activity of cellulo-somes was reported to be higher than, or at leastcomparable to, that of the fungal cellulases on purecellulose substrates (30).

Despite the different mechanisms of fungalcellulases and cellulosomes revealed in this study,cell wall materials are completely digestible byeither when lignins are effectively removed. Ther-mochemical pretreatment strategies to enhancebiomass digestibility by partial removal or re-distribution of lignin have been developed (31),however, these mechanisms result in sugar deg-radation and loss at high severities (32). Thechallenge now is to effectively and economical-ly modify lignins via strategies that maintainthe integrity of fermentable sugars. Researchershave recently focused on genetically engineer-ing plants for desirable lignin contents or com-positions that are more amenable to classicalpretreatment at low severities (33–35). In theforeseeable future, we expect that lignins in ge-netically modified energy plants will be extracted

under mild conditions and used as valuable chem-icals; the remaining cell wall architecture couldbe left intact with minimum alteration of thepolysaccharides. In this scenario, effective diges-tion by enzymes, especially fungal cellulases,could provide near-theoretical yields of ferment-able sugars.

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31. B. S. Donohoe et al., Biotechnol. Bioeng. 103, 480(2009).

Fig. 4. Size, shape, and arrangement of the CEFs. (A) An individual twisted CEFshown in delignified pSW surfaces. Composite-average profile of line scansacross the CEF appear in (B) vertical orientation [red dotted-line in (A)] and (C)horizontal orientation [green dotted-line in (A)]. (D) An untreated pSW showingmacrofibrils and microfibrils. (E) Composite-average profile of line scans acrossmacrofibrils [white dotted-line in (D)] containing two CEFs. Data are mean T SDbased on measurements of five well-defined microfibrils (B and C) andmacrofibrils (E). (F) Width measurements of 50 well-resolved fibrils (left) andaverage fibril-fibril distances measured in the entire image (right). Ten im-ages were measured for untreated and delignified PWs, pSWs, and sSWs.Macrofibril widths containing two to five CEFs in a PW were determined directlyby tracing fibril splitting. Horizontal bars denote mean values. (Inset) Dimensionsof the 36-chain CEF model are 5.3 nm (horizontal, green arrow) by 3.2 nm

(vertical, red arrow). The hydrophobic CEF faces are indicated in red. Furthermore,one CEF in the PW is expected to display the horizontal orientation, which pro-vides an apparent width of 5.3 nm based on cellulose structure. After delignifi-cation in pSWs, the vertical orientation is expected, which provides an apparentwidth of 3.2 nm. (G) Schematic illustration showing assembly of main cell wallcomponents in the PW and the sandwich-like SW. Scale is based on the averagefibril-fibril distance measured in (F). Cell wall orientations are indicated as: (a)transverse and (b) tangential relative to the cell lumen, and (c) cell long axis.Color bars in (A) and (D) represent the scale of the AFM height images; thewhite arrow in (A) indicates macrofibril splitting. Scale bar: 10 nm (A, D, and G).Additional images showing macrofibrils and microfibrils in untreated anddelignified PWs, pSWs, and sSWs are presented in fig. S5 to S7, respectively. Theproposed macrofibril splitting and rotating process is shown in fig. S8.

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32. R. Kumar, C. E. Wyman, Biotechnol. Bioeng. 102, 1544(2009).

33. C. X. Fu et al., Proc. Natl. Acad. Sci. U.S.A. 108, 3803(2011).

34. F. Chen, R. A. Dixon, Nat. Biotechnol. 25, 759 (2007).35. F. Chen, Y. Tobimatsu, D. Havkin-Frenkel, R. A. Dixon,

J. Ralph, Proc. Natl. Acad. Sci. U.S.A. 109, 1772(2012).

Acknowledgments: We thank M. P. Tucker for the fungalcellulases; S. Xie for guidance regarding SRS microscopy;K. Ruckman for manuscript editing; and A. J. Ragauskas,

C. E. Wyman, M. F. Davis, D. J. Johnson, and R. H. Atallafor valuable discussion. This work was supported by the U.S.Department of Energy (DOE) under contract no. DE-AC36-08-GO28308 with the National Renewable Energy Laboratory.We acknowledge research support from the BioEnergy ScienceCenter, a DOE Bioenergy Research Center, and the GenomicScience Program (ER65258), both supported by the Officeof Biological and Environmental Research in the DOE Officeof Science. S.-Y.D. conceptualized the project, conductedAFM, analyzed the data, and wrote the manuscript. Y.-S.L.conducted enzyme labeling, bright-field light microscopy, andCLSM. Y.Z. conducted SRS microscopy. E.A.B. purified the

cellulosomes. S.-Y.D., M.E.H., J.O.B., and E.A.B. revised themanuscript. We declare no competing financial interests.

Supplementary Materialswww.sciencemag.org/cgi/content/full/338/6110/1055/DC1Materials and MethodsFigs. S1 to S8References (36–40)Movies S1 to S4

16 July 2012; accepted 21 September 201210.1126/science.1227491

Quantum-State Resolved BimolecularCollisions of Velocity-ControlledOH with NO RadicalsMoritz Kirste,1* Xingan Wang,1* H. Christian Schewe,1 Gerard Meijer,1 Kopin Liu,2

Ad van der Avoird,3 Liesbeth M. C. Janssen,3 Koos B. Gubbels,3

Gerrit C. Groenenboom,3† Sebastiaan Y. T. van de Meerakker3,1†

Whereas atom-molecule collisions have been studied with complete quantum-state resolution,interactions between two state-selected molecules have proven much harder to probe. Here,we report the measurement of state-resolved inelastic scattering cross sections for collisions betweentwo open-shell molecules that are both prepared in a single quantum state. Stark-deceleratedhydroxyl (OH) radicals were scattered with hexapole-focused nitric oxide (NO) radicals in acrossed-beam configuration. Rotationally and spin-orbit inelastic scattering cross sections weremeasured on an absolute scale for collision energies between 70 and 300 cm−1. These cross sectionsshow fair agreement with quantum coupled-channels calculations using a set of coupled modelpotential energy surfaces based on ab initio calculations for the long-range nonadiabatic interactionsand a simplistic short-range interaction. This comparison reveals the crucial role of electrostaticforces in complex molecular collision processes.

Rotationally inelastic scattering is one ofthe key processes underlying the exchangeof energy between molecules (1, 2). In

bulk systems, rotational energy transfer (RET) isresponsible for the thermalization of state pop-ulations after a chemical reaction. In the diluteinterstellar medium, inelastic collisions contributeto the formation of nonthermal population dis-tributions that result in, for instance, interstellarmasers (3). Accurate state-to-state inelastic scatter-ing cross sections are essential ingredients for re-liablemodels of chemical processes in combustionphysics, atmospheric science, and astrochemistry.

In molecular beam collision experiments, theability to prepare molecules in a single rotational(sub)level before the collision using electric, mag-netic, or optical fields has been imperative tounravel the underlying mechanisms of molecularenergy transfer. This has made scattering exper-

iments possible at the full state-to-state level andhas resulted in the discovery of propensity rulesfor inelastic scattering (4), the stereodynamics ofmolecular collisions (5, 6), and quantum inter-ference effects (7–9). The latest beam decelera-tion and acceleration methods (10, 11) allow forthe precise variation of the collision energy, re-sulting in the observation of quantum thresholdeffects in the state-to-state cross sections (12, 13).This wealth of studies has contributed enormous-ly to our present understanding of how inter-molecular potentials govern molecular collisiondynamics.

Thus far, these methods have mostly beenused to study collisions of state-selected mole-cules with rare gas atoms. Yet, in most naturalenvironments,molecule-molecule interactions playa major role. For instance, space telescope ob-servations of cometary water may reveal the pos-sible origin of water on Earth, but a conclusiveinterpretation requires accurate knowledge of RETin water-water collisions (14). Whereas atom-molecule scattering cross sections can now becalculated routinely in excellent agreement withexperiment (13, 15), much less is known aboutRET in molecule-molecule collisions (16). Asopposed to an atomic target, a molecular scat-tering partner possesses internal degrees of free-dom of its own, adding a level of complexity that

can easily render ab initio quantum scatteringcalculations extremely challenging, if not im-possible. This is particularly true for collisionsinvolving radical species that are governed by mul-tiple Born-Oppenheimer (BO) potential energysurfaces (PESs) with nonadiabatic couplings be-tween them. Experimental data on bimolecularstate-to-state cross sections is generally lacking,and kinetic models often use collision rate coef-ficients that are expected to be inaccurate (17).

The study of molecule-molecule collisions atthe ultimate quantum level has been a quest inmolecular beam physics since it was establishedin the 1950s (18). Major obstacles exist that haveprevented studies of state-to-state bimolecularscattering (19). The main challenge is the needfor reagent beams with sufficient quantum-statepurity at the densities necessary to observe pop-ulation transfer in one, or both, reagent beam(s).Thus far, experiments of this kind have only beenpossible using cryogenically cooled H2 mole-cules as a target beam (20, 21).

Here, we report the successful measurementof state-resolved inelastic scattering between twostate-selected molecular beams. We have chosenthe OH (X 2P) + NO (X 2P) system (22) for thispurpose, as both open-shell radical species arebenchmark systems for the scattering of state-selected molecules with rare gas atoms that in-volve two BO PESs (23). Collisions between OHand NO involve eight interacting PESs, repre-senting the full complexity of bimolecular in-elastic collisions (24). The OH-NO system servesalso as a prototypical example of radical-radicalreactions of fundamental importance in gas-phasechemical kinetics (25). We used a Stark deceler-ator and a hexapole state selector in a crossed mo-lecular beam configuration to produce reagentbeams of OH and NO radicals with an almostperfect quantum-state purity. The collision ener-gywas varied between 70 and 300 cm−1 by tuningthe velocity of the OH radicals before the collisionusing the Stark decelerator, revealing the quantumthreshold behavior of the state-to-state inelasticscattering cross sections. The unusuallywell-defineddistributions of reagent molecules allowed us todetermine absolute scattering cross sections, whichcan normally be determined only on a relative scalein crossed-beam experiments. These cross sectionsshowed fair agreement with a theoretical modelfor inelastic collisions between two 2P radical spe-cies, based solely on an accurate description ofthe full rotational and open-shell structure of both

1Fritz-Haber-Institut der Max-Planck-Gesellschaft, Faradayweg4-6, 14195 Berlin, Germany. 2Institute of Atomic and Mo-lecular Sciences (IAMS), Academia Sinica, Taipei, Taiwan 10617.3Radboud University Nijmegen, Institute for Molecules andMaterials, Heijendaalseweg 135, 6525 AJ Nijmegen, TheNetherlands.

*These authors contributed equally to this work.†To whom correspondence should be addressed. E-mail:basvdm@science.ru.nl (S.Y.T.v.d.M.), gerritg@theochem.ru.nl (G.C.G.)

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