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Fast Intracrystalline Hydration of β-Chitin Revealed by Combined Microdrop Generation and On-Line Synchrotron Radiation Microdiffraction Manfred Ro¨ssle, David Flot, Jens Engel, Manfred Burghammer, and Christian Riekel* European Synchrotron Radiation Facility, B.P. 220, F-38043 Grenoble Cedex France
Henri Chanzy Centre de Recherches sur les Macromolecules Ve´ ge´ tales, CNRS, affiliated with the Joseph Fourier University of Grenoble, B.P. 53X, 38041 Grenoble Cedex9, France Received January 18, 2003; Revised Manuscript Received March 18, 2003
Water microdrops of about 50 µm in diameter, generated by an ink-jet system, have been used to hydrate fragments of Pogonophora tubes. In situ X-ray microdiffraction with a beam size of 10 µm was used to follow the structural transformations that affected the crystalline β-chitin part of the specimens. Starting from anhydrous chitin, the formation of a full β-chitin dihydrate was observed within about 90 s. A disordered intermediate phase with variable d-spacing that could be due to a mixture of anhydrous and hydrated β-chitin layers was also detected. Introduction Fluid quantities down to the picoliter (pL) range can be dispensed with a high temporal and positional precision using the ink-jet technology.1 At flight velocities of about 2 m/s, it is possible to increase the liquid concentration at a specific point of a solid on the subms time scale. For droplets of sufficiently small dimensions (water: e100 µm), the surface energy becomes larger than the kinetic energy, and the droplets merge with the solid without splashing. For a subsequent liquid/solid reaction, one can in this way very precisely initiate the reaction spatially and temporally. The aim of this report is to show that synchrotron radiation (SR) X-ray microdiffraction (µXRD) with beam sizes in the µm range2 is a convenient tool for investigating the kinetics of microstructural changes induced by liquid microdrops in a solid. As a demonstration example, we have investigated the intracrystalline hydration of β-chitin, which is known to form several hydrates.3-6 For specimen, we have chosen the highly crystalline β-chitin occurring in tubes from Birsteinia, a member of the Pogonophora phylum.7 Experimental Section Microdrop Generation. A commercial ink-jet system (Microdrop, Norderstedt, Germany) was used. The dispensing head contains a glass capillary mounted concentrically in a piezoelectric actuator. The system can be used for liquids with a viscosity in the range between 0.5 and 20 mPas. Distilled water was used for the hydration process. The diameter of the microdrops, which is defined by the capillary exit diameter, is about 50 µm corresponding to about 65 pL. * To whom correspondence should be addressed.
The relative change in volume of consecutive microdrops is specified by the manufacturer to be (1%. The control unit of the ink-jet system was triggered by a TTL signal from a VME frequency generator board. Software control was possible through a SPEC interface (Certified Scientific Software). X-ray Diffraction Setup. The dispensing head and the liquid reservoir were adapted to the microgoniometer of the ID13 beamline at the European Synchrotron Radiation Facility (ESRF; Figure 1, parts A and B).8,9 The capillary exit of the dispensing head was located about 200 µm above the sample position. Experiments were performed at RT. The microdrops were observed with a videomicroscope consisting of a microscope objective (coaxial to the X-ray beam), a 45° mirror, and a CCD camera at 90° angle. Stroboscopic light for sample observation was emitted by a photodiode (also coaxial to the X-ray beam), which was integrated in the illumination system of the microgoniometer. The X-ray optical system of the ID13 beamline has been described elsewhere.2 A monochromatic X-ray beam (λ ) 0.1008 nm) was focused through a bore in the objective/ mirror assembly to the sample position by an upstream condensing mirror. A Pt/Rh aperture at 20 mm from the center of the goniometer defined the beam size at the sample position to about 10 µm. The path of the microdrops was first located with the videomicroscope. The X-ray beam was then steered horizontally to match with the microdrop’s vertical trace. A specific point on the sample was then selected and placed in the beam position. Consecutive µXRD patterns (frames) were recorded with a slow-scan CCD camera (MAR) with 2048 × 2048 pixels, 64.5 × 64.5 µm2 pixel size and 16 bit readout. The full-frame readout time of about 10 s was reduced to about 1.5 s by binning the detector to 512 × 512
10.1021/bm0340218 CCC: $25.00 © 2003 American Chemical Society Published on Web 04/22/2003
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Figure 2. Image of Birstenia tube fragment aligned on microgoniometer. An arrow indicates the tube axis direction. The size of the X-ray beam is indicated by the white spots. µXRD patterns (frames) were recorded sequentially at the 5 spots during hydration (see below). A larger gray spot above the tube fragment shows the size of a 50 µm diameter water microdrop.
Figure 1. (A) Schematic setup of the microdrop generating system installed on the microgoniometer. (B) Image of the setup prior to starting experiment: a, microscope objective coaxial to X-ray beam direction; b, rotation axis; c, sample pin on sample holder; d, dispensing head; e, diode-light support; f, liquid reservoir.
pixels. The distance of sample to detector was determined to be 247.8 mm by an Ag-behenate standard.10 β-Chitin Sample. Dried Birsteinia tubes were kindly provided by the late K. M. Rudall. These tubes, which are in vivo about 0.5 mm in diameter and few centimeters in length, are synthesized by small deep-sea marine worms and served as their habitat. The tube wall consists of arrays of parallel chitin microfibrils, packed into flat ribbons, which are themselves superimposed in a crisscross manner, each ribbon making a distinct angle with respect to its underlying neighbors. Within the tube walls, the chitin microfibrils are embedded into a protein matrix in the ratio of roughly two parts of protein to one part of chitin.11 The dried tube had an oval cross section (large diameter ≈1.0 mm, small diameter ≈0.3 mm). Tube fragments were cut and glued on glass tips, which were fixed to standard sample holders (Hampton Research). The samples were then dried for a few hours in a vacuum oven at 50 °C. Prior to a hydration sequence, the sample was kept in air for about 15 min during transfer and alignment on the microgoniometer. Results Anhydrous β-Chitin. An optical image of the tube fragment after loading on the microgoniometer rotation axis
is shown in Figure 2. An arrow indicates the direction of the tube axis. The beam position is about 20 µm from the top surface of the tube as indicated by the white spots, which correspond to beam positions during data collection (see below). The frame shown in Figure 3A was recorded in 0.1 s. It corresponds essentially to the pattern of anhydrous β-chitin.3-6,12 We did not observe additional reflections or short-range order signal due to protein up to the edge of the pattern (Qmax ≈ 16 nm-1). The comparison of reflection intensities on both sides of the meridian suggests fiber symmetry. We find a bimodal distribution of the 010reflection azimuthal width of 12° fwhm (full-width halfmaximum; about 10%) and 69° fwhm. (Figure 3B) An additional weak equatorial 010-reflection13 of β-chitin monohydrate3 is probably due either to the exposure of the sample to air prior to the experiment or to imperfect dehydration (see above). The monohydrate reflection shows the same bimodal distribution as the anhydrous phase reflection. It is likely that any unwanted rehydration at this stage could be avoided by using a dry air stream from a cryoflow system. The radial intensity distribution in the range of the 010reflection integrated along the full azimuthal range is shown in Figure 3C. We used the FIT2D program14 for fitting the intensity distribution with an analytical function. We find that Gaussian functions can fit the intensity distribution significantly better than Lorentzian or Voigtian functions. The experimental profile is best matched by assuming that the anhydrous β-chitin reflection is composed out of two peaks representing two fractions of different particle sizes. The volume fractions can be derived from the relative integrated peak intensities. Based on the Scherrer formula,15 the major fraction (≈70 volume %) corresponds to an apparent particle size in the b direction of 18 nm and the minor fraction to 9 nm. These two fractions were found also at other positions on the azimuth and seem therefore to be independent from the bimodal distribution. (Figure 3B) We also find that a bimodal distribution is maintained throughout the hydration process (see below), which suggests morpho-
Fast β-Chitin Hydration
Figure 3. (A) µXRD pattern of the Birstenia tube fragment recorded in 0.1 s with a 10 µm diameter beam. The vertical arrow indicates the orientation of the tube-axis. The pattern corresponds mainly to the anhydrous β-chitin structure.3-6,12 Selected β-chitin reflections and the 010 reflection of the β-chitin monohydrate (M010) are indicated. (B) Radially averaged azimuthal intensity distribution of β-chitin 010 reflection before hydration. Two Gaussian functions and a flat background have been fitted to the profile. (C) Azimuthally (full angular range) averaged µXRD pattern in the range of the 010 reflections. Three Gaussian functions and a flat background have been fitted to the profile using the FIT2D program.14 The residuals correspond to the difference between the experimental curve (crosses) and the fitted profile (solid line).
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logical differences in the sample along the beam path. The corresponding particle size for the β-chitin monohydrate is 8 nm. In previous studies with vestimentiferan tube chitin, it was noted that transmission electron microscopy on tubes of TeVnia jerichonana worms shows that the major fraction of β-chitin microfibrils have somewhat larger diameters of 30-50 nm as compared to the present Birsteinia major fraction.16 Small microfibrils with 10 nm diameter microfibrils in T. jerichonana correspond, however, closely to those of the Birsteinia minor fraction. As discussed below, the drying of β-chitin microfibrils is also known to induce specific fracturing that could account for the fraction of smaller crystals.16 Intracrystalline Hydration. The hydration reaction was initiated by a 10 Hz microdrop sequence (t ) 0 s). During the reaction, the sample was displaced in a repetitive way through 40 µm by 5 steps parallel to the surface in order to reduce local radiation damage (Figure 2B). After every step, a 0.1 s frame was recorded. The formation of hydrated β-chitin can be readily observed after a few seconds by the appearance of an additional powder ring (Figure 4A). Maximum hydration is essentially complete in less than 2 min, which corresponds to an accumulated water volume of about 100 nL. For further analysis, the frames were azimuthally integrated and the 1D radial intensity profiles fitted with Gaussian profiles as shown in Figure 4B. Based on values reported in the literature, the positions of the 010 reflections of anhydrous β-chitin (A: 0.917 nm),3,4 monohydrate (M: 1.04 nm),3 and dihydrate (D: 1.128 nm)5 are indicated. The 010-reflection width and hence particle size of anhydrous β-chitin (minor fraction) and the β-chitin dihydrate are very similar. The position of the weak monohydrate reflection is only at the expected value for the t ) 0 s frame. As the reaction proceeds, we observe, however, that the position of this reflection varies gradually from the anhydrous phase to the dihydrate phase reflection. The value of d ) 1.101 nm for the dihydrate reflection is slightly smaller than the value reported in the literature (1.128 nm).5 The temporal evolution of phases can be seen more clearly in Figure 5 where the variation of the radial intensity distribution during the hydration process is shown. The reaction apparently does not pass through a stable intermediate monohydrate phase with the b-axis parameter reported in the literature3. We note a slight increase of the background level toward smaller Q values upon hydration, which requires the use of a first order polynomial in the fitting procedure. Kinetics. For an analysis of the hydration kinetics, the 010 profiles of anhydrous β-chitin and dihydrate were azimuthally integrated in a limited Q range around the ideal position and fitted by a Gaussian function. The temporal variation of the integrated intensities is shown in Figure 6. Intensity values have been corrected for beam intensity decay and scaled to 1.0 at maximum intensity of each curve. Monohydrate intensity values could not be derived in the same way as the peak position of the intermediate phase changes. For anhydrous β-chitin, we find after an incubation period a practically linear intensity decrease with a half time τ1/2 ≈ 65 s. The dihydrate phase starts obviously to form at the onset of the hydration process. We note that the intensity
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Figure 4. (A) Selected frames recorded during the hydration sequence with 10 Hz water microdrops. The Q range has been restricted to the 010 reflection. t ) 0 s corresponds to the onset of microdrop generation. (B) azimuthally (full angular range) integrated patterns. The position of the 010-reflections of anhydrous β-chitin (A), monohydrate (M), and dihydrate (D) phases based on values from literature (see text) are indicated. At 93.2 s, a mixture of the anhydrous and the two hydrated β-chitin phases is observed, whereas mostly dihydrate β-chitin is present at 223.6 s (see also text).
of the 010 reflections of anhydrous β-chitin and dihydrate match at a scaled relative intensity of about 0.3 (t ≈ 80 s). At longer reaction times (t > 250 s), we find a decrease of dihydrate 010-reflection intensity, which is probably due to radiation damage. Discussion Time-resolved synchrotron radiation studies on the hydration of biopolymers such as DNA17 are well-known. The results described in this study are, however, novel as one was able to follow the penetration of water molecules within the crystalline domains of a biopolymer at a precise location and with an accurate timing. The striking feature revealed when following the kinetics of intracrystalline hydration of β-chitin is that it takes less than 2 min to convert a nearly pure anhydrous phase into a full dihydrate that denotes a lattice expansion of more than
0.2 nm in the b-axis direction. Another aspect revealed by the present study is that the β-chitin monohydrate phase does not appear as a stable intermediate phase, at least in the fast hydration system where drops of liquid water hit the sample surface. In fact, a figure such as the one shown in Figure 5 tends to show that there are only two stable crystallographic phases in the β-chitin system: the anhydrous and the dihydrate phases. The observation of two sizes of crystals in anhydrous or hydrated in Birstenia specimens may be explained in at least two ways in relation with the substantial ultrastructural work achieved with chitin from vestimentiferan worms,16,18,19 which shows a close similarity with the chitin from Pogonophora. At first, one can consider that there are indeed different sizes of crystals that are produced by the worms when they synthesize the tubes. Indeed, ultrathin sections perpendicular to the chitin microfibrils reveal a distribution of lateral sizes of the microfibrils, even within a same ribbon
Fast β-Chitin Hydration
Figure 5. Temporal variation of the azimuthally averaged (010) reflection profiles (Figure 4B). The intensity (expressed in false color) is logarithmically displayed. The d values of the different phases (A/ M/D) correspond to values reported in the literature (see text). The relative height of the peaks at 0/93.2/223.6 s can be obtained from Figure 4B (see also Figure 6).
Figure 6. Temporal variation of integrated intensity the 010 reflections of anhydrous β-chitin (A) and β-chitin dihydrate (D). The intensity at t ) 0 was scaled to the average values of the 8 frames preceding the hydration onset which was set to 1. The dihydrate 010 intensity was scaled to the maximum observed intensity ()1).
containing an array of parallel microfibrils.11,16,18 It is likely that the origin of such a distribution can be found in the various sizes of the organelles that synthesize chitin microfibrils within the chitin-secreting glands. A second possibility would consider the fact that the larger chitin crystals were broken when the sample that we used was dried and stored prior to the experiment. Indeed, as known from the parent work on vestimentiferan chitin, the chitin crystals with diameter as large as 50 nm are biosynthesized under dihydrate conditions. Upon drying, these crystals shrink by about 25% in the b direction. This shrinkage induces a number of fractures that are clearly observed in the cross sections,16 with the result of a distribution of smaller crystalline domains that could fall in two categories: the small and large fragments that are deduced from the deconvolution shown in Figure 3. At present, it is not
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possible to distinguish between these two hypotheses, but planned future work with once dried and never dried specimens should indicate whether the diminution in crystal size is consistent with artifactual defects resulting from the drying. As previously mentioned, the ultrastructure of Birstenia tubes appears to be similar to that of vestimentiferan tubes and in particular to that of T. jerichonana, which consists of a superposition of chitin ribbons where arrays of β-chitin microfibrils are embedded in an amorphous protein matrix.16,18-20 The swelling of β-chitin with water is known to be strongly anisotropic without modification of the β-chitin sheets that are maintained intact by the strong NH‚‚‚OdC intermolecular hydrogen bonds.4,6 An analogue process can be found in intercalation reactions of sheet silicates,21 graphite,22 or transition metal dichalcogenide23 where the transition between two phases (stages) can be modeled by stacking disorder. Such intermediate phases can show a variation in peak position and a broadening due to a change in stacking disorder, which is also observed in the present case. This suggests that the intermediate phase crystallites contain a random mixture of anhydrous and hydrated β-chitin “sheets” in varying proportions as the hydration proceeds. A slight admixture of anhydrous phase layers might also be present for the dihydrate phase as the observed d spacing is slightly smaller than the value expected from literature (see above). One may wonder why a transient β-chitin monohydrate step was not observed during our on-line hydration experiment. In fact, our observations indicate that even the small initial fraction of this phase quickly disappears as soon as the first water drops hit the sample. Thus, it seems that, if enough water is available, the stable phase will always be the β-chitin dihydrate, which corresponds to the initial native state of β-chitin in its marine environment.16 The absence of a monohydrate phase in our experiment agrees well with the study of Saito et al.,6 who have shown that in a close aqueous environment a substantial monohydrate phase was observed only when the β-chitin samples were heated above 100 °C. Discussing DSC and X-ray diffraction results of the hydration/dehydration cycles of their β-chitin specimens, these authors could conclude that the conversion of anhydrous β-chitin to monohydrate was a slow process as opposed to the conversion of monohydrate to the dihydrate which was fast. Thus, if an excess of water is present and if the sample is kept at room temperature, one should never see any monohydrate, in full agreement with what we are observing in the present experiments. Our diffraction data differ, however, from those of Saito et al.6 in the sense that we are following on-line the onset of the dynamics of hydration, whereas their study corresponded rather to an equilibrium state. In this respect, Figure 6 brings some new feature as it clearly shows that the disappearance of the anhydrous phase reflection occurs at a faster rate than the appearance of the dihydrate phase reflection, but nevertheless without showing any hint of transient monohydrate phase. Details of the reaction mechanism will have to be worked out in future studies but the slow onset of decay of the anhydrous phase suggests a nucleation and growth type
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kinetics. It seems that nuclei growing at the surface of the β-chitin crystallites transform these crystallites at a critical size quickly into the disordered, intermediate phase. It is only in a slower subsequent step that the water molecules become structured into the lattice to yield the ordered dihydrate phase. Thus, if the rate of formation of the dihydrate phase is fast compared to that of the monohydrate, it is nevertheless slower than the rate of the intermediate phase formation. In the world of crystalline neutral polysaccharides, most specimens occur under different forms of crystalline hydrates. In these crystals, the water molecules adopt either a sheetlike structure as in the present or are located within columns.24 Similar to β-chitin is the orthorhombic nigeran, where again the incorporation of water inside the anhydrous lattice results in an expansion of 0.225 nm along only the b axis of the unit cell while the a and c (chain) axes remain nearly constant.25 In view of the similarity with the present observations, it is likely that the fast hydration kinetics of nigeran should be similar to that of β-chitin. Quite remarkably, the drying of nigeran crystals induces also spectacular cracks, similar to those occurring with β-chitin from vestimentiferan tubes. Samples where the intracrystalline water occurs as columns are quite abundant. These crystalline hydrates are found in the A26 and B27 amylose that constitute the crystalline domains of starch, (1 f 3)-β-D28 and (1 f 4)-β-D29 xylans, polysaccharides of the (1 f 3)-β-D glucan family,30 etc. For all of these samples, no kinetics data exists that indicates the rate of penetration of water in the crystals, within interchain column. Experiments such as the one presented here achieved with either starch granules or fibers of (1 f 3)-β-D glucans can be envisaged. The resulting data should bring some interesting view on the columnar hydration dynamics. Indeed, whether the sheetlike hydration of β-chitin is likely to progress readily in a wedge-like fashion between chitin sheets, a more complicated system must occur in the columnar case. A much slower kinetics is expected for the diffusion of columnar hydration, as the water must percolate from columns to columns throughout the crystalline domains. This remains to be demonstrated experimentally with the present setup. Set aside hydration processes in solids the ink-jet technology in combination with µXRD or micro-X-ray small-angle scattering (µSAXS)31 could find multiple other applications including fast mixing processes of microdrops. Acknowledgment. J. Meyer and H. Gonzalez (ESRF) interfaced the ink-jet system to the beamline control system. L. Lardiere developed the mechanical support structure.
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References and Notes (1) (2) (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13)
(14) (15) (16) (17) (18) (19)
(20)
(21) (22) (23) (24) (25) (26) (27) (28) (29) (30) (31)
Lee, E. R. In Microdrop Generation; CRC Press: New York, 2002. Riekel, C. Rep. Prog. Phys. 2000, 63, 233. Blackwell, J. Biopolymers 1969, 7, 281. Saito, Y.; Okano, T.; Gaill, F.; Chanzy, H.; Putaux, J.-L. Int. J. Biol. Macromol. 2000, 28, 81. Saito, Y.; Okano, T.; Putaux, J.-L.; Gaill, F.; Chanzy, H. In AdVances in Chitin Science, Volume II; Domard, A., Roberts, G. A. F., Vårum, K. M., Eds.; Jacques Andre´ Publ.: Lyon, France, 1998; p 507. Saito, Y.; Kumagai, H.; Wada, M.; Kuga, S. Biomacromolecules 2002, 3, 407. Blackwell, J.; Parker, K. D.; Rudall, K. M. J. Mar. Biol. Assn. U.K. 1965, 45, 659. Cusack, S.; Belrhali, H.; Bram, A.; Burghammer, M.; Perrakis, A.; Riekel, C. Nature Struct. Biol. 1998, 5, 634. Perrakis, A.; Cipriani, F.; Castagna, J. C.; Claustre, L.; Burghammer, M.; Riekel, C.; Cusack, S. Acta Crystallogr. 1999, D55, 1765. Blanton, T. N.; Huang, T. C.; Toraya, H.; Hubbard, C. R.; Robie, S. B.; Louer, D.; Go¨bel, H. E.; Will, G.; Gilles, R.; Raftery, T. Powder Diff. 1995, 10, 91. Gupta, B. L.; Little, C. Z. Zool. Syst. EVolutionsforsch. 1975, 13, 45. Gardner, K. H.; Blackwell, J. Biopolymers 1975, 14, 1581. In this work, we are using for anhydrous β-chitin the unit cell defined as in ref 3 (a ) 0.485 nm, b ) 0.926 nm, c (fiber axis) ) 1.023 nm, and γ ) 97.5°). For chitin dihydrate, we use the parameters defined as in ref 5 (a ) 0.489 nm, b ) 1.128 nm, c ) 1.038 nm, and γ ) 96.8°). Web page: /www.esrf.fr/computing/scientific/FIT2D/FIT2D_REF/ fit2d_r.htm. Klug, H. P.; Alexander L. E. In X-ray Diffraction Procedures for Polycrystalline and Amorphous Materials, 2nd ed.; Wiley-Interscience: New York, 1974; p 687. Gaill, F.; Persson, J.; Sugiyama, J.; Vuong, R.; Chanzy, H. J. Struct. Biol. 1992, 109, 116. Fuller, W.; Mahendrasingam, A.; Denny; R. C.; Forsyth, V. T. Pigram, W. J.: Papiz, M. Proc. Ital. Phys. Soc. 1990, 25, 991. Tanner, S. F.; Chanzy, H.; Vincendon, M.; Roux, J.-C.; Gaill, F. Macromolecules 1990, 23, 3576. Gaill, F.; Schillito, B.; Chanzy, H.; Goffinet, G.; Da Conceicao, M.; Vuong, R. In AdVances in Chitin and Chitosan; Brine, C. J., Sandford, P. A., Zikakis, J. P., Eds.; Elsevier Applied Science: New York 1992; p 225. Gaill, F.; Persson, J.; Sugiyama, J.; Vuong, R.; Tanner, S.; Chanzy, H. In AdVances in Chitin and Chitosan; Brine, C. J., Sandford, P. A., Zikakis, J. P., Eds.; Elsevier Applied Science: New York 1992; p 216. Hendricks, S. B.; Teller, E. J. Chem. Phys. 1942, 10, 147. Metz, W.; Hohlwein, D. Carbon 1975, 13, 87. Riekel, C. Prog. Solid State Chem. 1980, 13, 89. Bluhm, T.; Deslandes, Y.; Marchessault, R. H.; Sundararajan, P. In Water in Polymers; ACS Symposium Series 127; American Chemical Society: Washington, DC, 1980; p 253. Taylor, K. J.; Chanzy, H.; Marchessault, R. H. J. Mol. Biol. 1975, 92, 165. Imberty, A.; Chanzy, H.; Perez, S.; Buleon, A.; Tran, V. J. Mol Biol. 1988, 201, 365. Imberty, A.; Perez, S. Biopolymers 1988, 27, 1205. Atkins, E. D. T.; Parker, K. D. J. Polym. Sci. Part C 1968, 28, 69. Nieduszynski, I.; Marchessault, R. H. Biopolymers 1972, 11, 1335. Chuah, C. T.; Sarko, A.; Deslandes, Y.; Marchessault, R. H. Macromolecules 1983, 16, 1375. Riekel, C.; Burghammer, M.; Mu¨ller, M. J. Appl. Crystallogr. 2000, 33, 421.
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