Cellulose (2009) 16:943–957 DOI 10.1007/s10570-009-9338-5
ORIGINAL PAPER
The structure of the complex of cellulose I with ethylenediamine by X-ray crystallography and cross-polarization/magic angle spinning 13 C nuclear magnetic resonance Masahisa Wada Æ Laurent Heux Æ Yoshiharu Nishiyama Æ Paul Langan
Received: 4 March 2009 / Accepted: 18 June 2009 / Published online: 11 July 2009 Springer Science+Business Media B.V. 2009
Abstract X-ray crystallographic and cross-polarization/magic angle spinning 13C nuclear magnetic resonance techniques have been used to study an ethylenediamine (EDA)-cellulose I complex, a transient structure in the cellulose I to cellulose IIII conversion. The crystal structure (space group P21; ˚ , b = 11.330 A ˚ , c = 10.368 A ˚ and a = 4.546 A c = 94.017) corresponds to a one-chain unit cell with one glucosyl residue in the asymmetric unit, a gt conformation for the hydroxymethyl group, and one EDA molecule per glucosyl residue. Unusually, there are no O–HO hydrogen bonds between the cellulose chains; the chains are arranged in hydrophobic stacks, stabilized by hydrogen bonds to the amine groups of bridging EDA molecules. This new structure is an example of a complex in which the
M. Wada Department of Biomaterials Science, Graduate School of Agricultural and Life Sciences, The University of Tokyo, Tokyo 113-8657, Japan L. Heux Y. Nishiyama Centre de Recherches sur les Macromole´cules Ve´ge´tales – CNRS, Affiliated with the Joseph Fourier University of Grenoble, BP 53, 38041 Grenoble Cedex 9, France P. Langan (&) Bioscience Division, Los Alamos National Laboratory, Los Alamos, NM 87545, USA e-mail:
[email protected]
cellulose chains are isolated from each other, and provides a number of insights into the structural pathway followed during the conversion of cellulose I to cellulose IIII through EDA treatment. Keywords X-ray crystallography NMR Cellulose Ethylenediamine
Introduction The use of amines has been studied and developed for many years now in the treatment and processing of cellulose and cellulosic biomass fibers (Clark and Parker 1937; Davis et al. 1943; Klemm et al. 1998; Klenkova 1967; da Silva Perez et al. 2003; Detroy et al. 1981; Umikalsom et al. 1997). They can act as swelling agents, penetrating fibers and forming crystalline complexes with cellulose, some of which can be relatively stable. On removal of amines, the fibers can have improved pulping, textile or biomass conversion properties. Ammonia treatments have been used to improve the strength and shrink resistance of cotton and regenerated fabrics (Yanai and Shimizu 2006; Pandey and Nair 1975; Lewin et al. 1974) and are being explored as pretreatments for biomass to improve its conversion to glucose and biofuels (Teymouri et al. 2005). Ethylenediamine (EDA) has also been investigated for its use in biomass pulping and pretreatment (Jahan and Farouqui 2000; Zargarian
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et al. 1988) and as a component of cellulose solvents (Xiao and Frey 2007). In order to understand the detailed interaction of amines with cellulose and cellulosic biomass fibers we have been studying these interactions using a variety of crystallographic and spectroscopic techniques. In previous studies it has been described in atomic detail how ammonia treatment transforms the naturally occurring forms of cellulose, Ia and Ib which are collectively called cellulose I (Sugiyama et al. 1990, 1991a, b; Nishiyama et al. 2002, 2003) into an intermediate ammonia-cellulose I complex (Wada et al. 2006) and then, on removal of ammonia by evaporation, into cellulose IIII (Wada et al. 2001, 2004), a crystal form of cellulose which is highly activated for hydrolysis by cellobiohydrolases (Igarashi et al. 2007). On hydrothermal treatment cellulose IIII is transformed back to cellulose Ib (Wada 2001). The Ia phase is considered to be less stable than the Ib phase because it can be irreversibly converted to Ib by hydrothermal treatment (Horii et al. 1987; Yamamoto et al. 1989). Thus we provided a structural pathway for the ammonia associated solid-state conversion of cellulose between different crystal phases. In a similar way, treatment with EDA can transform cellulose Ia or Ib into an intermediate EDAcellulose I complex, and then, on removal of EDA by washing in polar but non-aqueous solvents such as methanol or ethanol, into cellulose IIII (Chanzy et al. 1986, 1987). As in the case of ammonia treatment we can define a structural pathway for the solid-state conversion of cellulose between the different crystal phases associated with EDA treatment, Fig. 1. Detailed crystal structures for the cellulose Ia, Ib, IIII phases in this pathway have already been provided
Fig. 1 The solid-state conversion of cellulose between various crystal phases involving EDA and hydrothermal treatment
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(Nishiyama et al. 2002, 2003; Wada et al. 2001, 2004). A crystal structure has also been reported for EDA-cellulose I from X-ray diffraction studies (Lee et al. 1984), however recent cross-polarization/magic angle spinning (CP/MAS) 13C nuclear magnetic resonance (NMR) studies have raised doubts about the details of this structure (Numata et al. 2003). In early X-ray diffraction studies of cotton fibers treated with various diamines, a monoclinic twochain unit cell was reported for EDA-cellulose I ˚ , b = 12.3 A ˚ , c = 10.3 A ˚ , c = 43.2) (a = 12.2 A (Creely et al. 1959). More extensive X-ray diffraction data collected from ramie fibers treated with EDA allowed the determination of a more detailed structure; two parallel chains pack in a monoclinic unit ˚ , b = 9.52 A ˚, cell with P21 symmetry (a = 12.87 A ˚ c = 10.35 A, c = 118.8) (Lee et al. 1984). The two, almost identical, independent chains have the same translation along the chain (c-axis) direction and lie on 2-fold screw axes. There is one EDA molecule per glucosyl residue and therefore two EDA molecules and two glucosyl residues in the asymmetric unit. The hydroxymethyl groups of both chains are in the gt conformation. There are three low-energy conformations for the hydroxymethyl group in cellulose, designated tg, gg and gt; the first letter indicates whether the position of the O6 atom is either trans or gauche with respect to atom O5, and the second refers to its position with respect to atom O4. The general features of this X-ray model were confirmed by solid-state 13C CP/MAS NMR studies of an EDA-cellulose I complex prepared from Valonia cellulose (Chanzy et al. 1987; Henrissat et al. 1987). Single resonance lines for each carbon atom were observed, indicating the presence of only one glucosyl conformation, or at least near equivalent glucosyl conformations. The resonance lines for C1, C4, C6 and EDA were assigned, and the chemical shift of C6 at 62.2 ppm was consistent with a gt conformation for the hydroxymethyl group. More recent 13C CP/MAS NMR studies of EDAcellulose I, prepared from Valonia cellulose, also contained single resonance lines for each carbon (Numata et al. 2003). All carbon resonance lines were unambiguously assigned and their chemical shifts indicated that the conformation of the glucosyl residue differed significantly from that in the X-ray structure of EDA-cellulose I. However, a fitting analysis resulted in only one EDA molecule per two
Cellulose (2009) 16:943–957
glucosyl residues, in disagreement with the X-ray structure (Lee et al. 1984). We have developed improved methods for preparing highly crystalline fibers of EDA-cellulose I and we have collected high-resolution X-ray crystallographic data from those fibers (Wada et al. 2008). A preliminary analysis of the X-ray reflection positions in this data resulted in a new unit cell (space group ˚ , b = 11.33 A ˚ , c = 10.37 A ˚ and P21; a = 4.55 A c = 94.02) with a volume that is about half of that proposed in previous X-ray crystallographic studies (Lee et al. 1984). Although no crystal structure determination was carried out, on the basis of unit cell size, density measurements, and thermogravimetric analysis we were able to determine that this unit cell corresponds to one cellulose chain, with the asymmetric unit composed of one glucosyl residue and one EDA molecule. Although the presence of only one glucosyl residue in the asymmetric unit is in agreement with previous 13C CP/MAS NMR studies, the presence of one EDA molecule per glucosyl residue is not (Numata et al. 2003). In the present work we report the crystal structure of EDA-cellulose I based on a detailed analysis of the X-ray diffraction patterns we reported previously (Wada et al. 2008), and new 13C CP/MAS NMR spectra collected from those same samples. The new structure is significantly different from the previously reported X-ray structure (Lee et al. 1984) and provides new insights into the structural pathway followed during the solid-state conversion of cellulose by EDA. The structure also reveals similarities and differences in the detailed action of ammonia and EDA on cellulose. Our hope is that these structural explanations of the underlying actions of amines on cellulose, will prove useful for predicting new, or improved, cellulose and cellulosic biomass fiber amine treatments.
Experimental section Preparation of oriented cellulose I samples and X-ray data collection Sample preparation, conversion to EDA-cellulose I, and X-ray data collection have been described previously (Wada et al. 2008). Briefly, green algae Cladophora sp. were used in this study because
945
although the cellulose is a mixture of the Ia and Ib allomorphs it is highly crystalline. After purification (Sugiyama et al. 1991a, b), hydrolyzed cellulose microcrystals were reconstitution into oriented films (Nishiyama et al. 1997). Bundles of films were then immersed in anhydrous EDA, dried, and then synchrotron X-ray data were collected from them on beam line BL38B1 at the SPring-8. Solid state NMR data collection Samples were inserted individually into tightly sealed 4-mm BL type ZrO2 rotors. 13C CP/MAS NMR spectra were recorded with a Bruker Avance spectrometer equipped with a 4-mm BL type probe and operated at 100 MHz. The spectra were acquired at room temperature with a 100 kHz proton dipolar decoupling field, matched cross-polarization (CP) fields of 80 kHz, a proton 90 pulse of 2.5 ls and magic angle spinning (MAS) at a spinning speed of 12 kHz. The cross-polarization transfer was achieved using a ramped amplitude sequence (RAMP-CP) for an optimized total time of 2 ms. The sweep width was of 50,000 Hz to avoid baseline distortion with 2,994 TD points, and the Fourier transformation was achieved without apodization over 8 k points. The repetition time was 4 s and the average number of 20,000 scans was acquired for each spectrum. The 13 C chemical shifts were determined relative to the carbon chemical shifts of the carbonyl signal of glycine at 176.03 ppm. Figure 2 shows the spectra recorded from the cellulose I sample before treatment with EDA, the EDA-cellulose I sample, and the cellulose IIII sample after removal of EDA. The chemical shifts are reported in Table 1. X-ray data processing X-ray data were processed using the program suite FibreFix. The program XFIX in this suite was used to determine the image center and the values of the rotation and tilt of the sample. All of the Bragg reflections could be indexed on a lattice corresponding ˚, b = to a monoclinic unit cell with a = 4.546(5) A ˚ ˚ 11.330(5) A, c = 10.368(5) A and c = 94.017(5) as shown in Fig. 3. There were no systematic 00l absences, but meridional reflections on odd layer lines were relatively weak and therefore a space group of P21 was assigned.
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Fig. 2 13C CP/MAS NMR spectra recorded from a Cladophora sp. cellulose I (a mixture of Ia and Ib), b EDA-cellulose I, and c cellulose IIII
Table 1 The 13C chemical shifts recorded from the cellulose I sample before treatment with EDA, the EDA-cellulose I sample, and the cellulose IIII sample after removal of EDA Sample
C1
C2
105.7; 105.1; 104.2
ND
C3
C4
C5
C6
EDA
Chemical shifts (ppm) 89.9; 88.9; 88.1; 84.5a b
ND
65.3
EDA-cellulose I
105.2
71.7
77.5
84.6
76.5b
62.0
Cellulose III
104.9
73. 6b
76.0b
87.8
72.9b
62.3
Cellulose I
ND b
a
Disordered/surface chains
b
Assignments according to (Numata et al. 2003)
The detector image was then remapped into cylindrical reciprocal space using the program FTOREC. LSQINT was used to refine a two-dimensional fit of the background and Bragg intensities in reciprocal space. The background was fitted using the ‘‘roving aperture’’ method. A Lorentzian distribution was used to model the angular profile of the spots and the width of this distribution was refined together with the unit cell parameters during maximum entropy refinement. The relevant crystal data and summary of intensity data collection are given in Table 2. At the end of refinement, a standard deviation, r, was calculated for each reflection directly from the root-mean-square difference between the calculated and observed pixel intensity values over the predicted profile of the spot. All reflections with similar reciprocal radial distances were
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45.6
regarded as accidentally overlapping due to cylindrical averaging and the intensities associated with those reflections were merged. The measured and merged observed intensity values are given in Table 3. X-ray structure refinement X-ray structure refinement was carried out using previously described strategies for applying SHELX97 (Sheldrick 1997) to fiber diffraction data (Langan et al. 2001). Atomic starting positions were taken from the crystal structure of cellulose IIII (Wada et al. 2004). An initial refinement was carried out with the hydroxymethyl group O6 atom removed, with the chains arranged in the ‘‘parallel-down’’ configuration ˚ . The and using X-ray data out to a resolution of 1.5 A
Cellulose (2009) 16:943–957
947 Table 2 Crystallographic experimental details Chemical formula
Cellulose (C12H20O10)
EDA (C2H8N2) Crystal data Space group ˚) a (A
P21
˚) b (A ˚) c (A
11.330(5)
4.546(5) 10.368(5)
c () ˚ 3) V (A
94.017(5) 532.7
Data collection Independent refls
180
Reflections [ 2r(I)
164
hmax ()
26.2
Range of h
0 to 3
Range of k Range of l ˚) k (A
-7 to 7 0 to 8 1.00
Refinement Refinement on 2
2
F2
R[F [ 2r(F )]
0.164
Fig. 3 A detail of the synchrotron X-ray diffraction pattern collected from EDA-cellulose I. Predicted reflection positions ˚ , b = 11.330 A ˚, generated using a unit cell with a = 4.546 A ˚ and c = 94.017 and with space group P21 are c = 10.368 A shown in yellow. The fiber axis is vertical
xR(F2)
0.408
Dqmax
0.43
Dqmin
-0.56
qrms
0.12
‘‘parallel-up’’ and ‘‘parallel-down’’ structures are defined according to the definition of French and Howley (1989). Both positional parameters and individual thermal parameters were refined, involving a total of 124 reflections, 45 parameters with 43 applied constraints on those parameters. The resulting values of R and Rx were 0.3146 and 0.6688, respectively. R is calculated from R(|Fo|-|Fc|)/R|Fo| with Fo [ 4r, where Fo and Fc are the observed and calculated amplitudes respectively. Rx, is calculated from [Rx(F2o-F2c )2/Rx(F2o)2]1/2, where x is a weight (1/r2) applied to each F2 term in the least squares refinement. Rx will be more than twice the size of the R. The corresponding 2Fo-Fc and Fo-Fc Omit maps clearly indicated that the hydroxymethyl group was in the gt conformation (Fig. 4). Additional density in this map, that could not be assigned to cellulose atoms, indicated the positions of the EDA atoms. The EDA molecule was then fitted to the density in the Fo-Fc omit map, and a subsequent refinement, which also included the O6 atom in the gt position,
involved an additional 7 parameters and 6 constraints. The resulting values of R and Rx were 0.1681 and 0.4260, respectively. When the occupancy of the EDA molecule was included in the refinement, it refined to a value of 1.02 without significantly improving the values of R or Rx. The EDA occupancy was therefore fixed at a value of 1.0 in subsequent refinements. Refinement of all atoms using all of X-ray data ˚ increased the number of extending out about 1.2 A data to 200, did not change the structure greatly, and increased the values of R and Rx to 0.1920 and 0.5200. There was also a systematic deviation from a flat analysis of variance at high resolution, an effect previously reported in similar high resolution studies of cellulose (Langan et al. 2001). This effect is due to the intensity of weak reflections being overestimated at high angle where they have large extents due to arcing effects. LSQINT may fit weak reflections to background features that have been underestimated, and the background fitting method used in LSQINT
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Table 3 Measured observed intensity values, where h, k and l are Miller indices, I is intensity, rI is the error in intensity and m is the multiplicity of each reflection h k 0
l
rI
I
m
h k
l
1 0
37.25 3.36
0
2 0
11.9
2 1 -6 0
2.13 0.59
1
0 0 262.09 5.29
2 2
4 0
0
1 -1 0 312.88 4.91
2 1
6 0 14.02 0.66
1
1 0 245.91 5.6
2 2 -5 0
0
3 0
0
1.5
0
2 2 -4 0
rI
I
-2 0
7 0
1 -2 0
19.27 7.59
2 1 -7 0
1
2 0
19.55 2.02
2 2
1 -3 0
12.09 0.72
0
4 0
1
3 0
0
0
12.14 2.09
0
0 0
h k
l
I
rI
m
0
-2 1
2 0
5 1
0
0
-2 1 -1 2 12.09 0.7
-2 2 -1 1 52.91 1.03 2 2
1 1
0
0
3.32 0.25
2 1 -5 1
0
-2 2
0
2 1 14.33 1.19
2 2 -3 1
6.2
0.68
2 1 -2 0 -2 1
1 2 32.85 1.08 3 2
0
0
2 2 19.53 0.68
2 1 -3 2
0.93 0.62
2 0
4 2
0
2 1
3 2
9.57 0.48
0
-2 1
5 1 16.71 0.6
0
0
-2 2
3 1
0
2 0
6 1
4.78 0.27
2 1
0
0 2 53.12 1.24
2 1 -2 2 31.79 1.71
0
2 2 -6 0 14.82 1.09
rI
I
0
1.25
0
l
0 1
2 2 -2 1 60.6
0
h k
2 2
2.59 0.38
5 0 50.63 0.71
2 3 -1 0 -2 3
3.21 0.45
m
-2 1 -4 2
0
m 2 2 2 -2 2 2 2 -2 2
8.11 1.7
2
4 2
7.61 0.9
2
1 -4 0
24.26 0.77
2 0
1 1
0.58 0.06
2 2 -4 1
5.79 0.52
2 2
0 2
0
0
-2
1
4 0
14.39 2.82
2 0
2 1
2.44 0.52
2 1 -6 1
1.18 0.21
2 0
5 2
0
0
-2
2
0 0
2 2
0
0
0
-2 1
0 1 21.1
1.05
0 5 0 0 0 -2 1 -1 1 7.52 2.54 2 -1 0 107.25 3.12 2 1 1 1 12.42 2.23 2
1 0
2 -2 0 1 -5 0 2
0
0
19.93 2.86 0
0
2 0
12.91 2.15
2 -3 0
9.34 0.55
1
5 0
2
3 0
0
0
6 0
23.1
2
4 2
0
1
6 2
7.75 3.31 1.1
2 -5 2 0
7 2
37.5
0.9 0 1.09 0 0.27
2.65 0.28
-2 0
3 1
0
0
2 1 -2 1 29.84 2.79 -2 1
2 1 14.28 0.92
2 1 -3 1 13.39 0.64 2 0
4 1
0
0
2 1
3 1 36.42 1.7
-2 1 -4 1 21.54 0.49
4 1
0
2 1 6 1 12.57 0.42 2 2 -5 1 3.89 0.28 -2 0
7 1
2 1 -7 1 2 2
0
0
5 1 18.09 0.28
2 3 -1 1 -2 3
1.85 0.27
0 1
2 2 -6 1
2 1 -5 2 -2 2
0
0
2 -2 2 -2
2 2 16.28 1.25
2
2 2 -3 2 11.58 0.35
2
-2 1
5 2 35.89 0.44 3 2
4.3
0.44
2 0
0
0
6 2 12.62 0.44
2 -2 2
2 0
1 2 26.41 0.89
2 2 -4 2
2.4
0.18
2
4 1 53.57 1.56
2 0
2 2 50.15 1.17
2 1 -6 2
2
0.22
2
6 3
2 2 -3 4
6.98 0.31
2 2 -2 5 21.6
0.82
2 2 -4 3 14.78 0.39
2 1
5 4
4.28 0.31
2 1 -5 5
0
2 1 -6 3
2 2
3 4
0
-2 0
6 4
8.41 0.31
2 1 -2 0
4.81 0.38 2.13 0.27
2 5 2 3 -1 2
7.98 0.36 2 2 -5 3 0 0 -2 0 7 3
2.46 0.42
2 1 -6 4
5.39 0.46
2.74 2.84
2 2
4 4
0
3
0
0
-2 1
6 4
4.94 0.31
-2 1 -7 3
2 -6 2
14.41 0.47
2 2
0
32.22 1.14
2 3 -1 3
5 3
0
2.46 0.18
2 2 -4 4 17.52 0.5
2 2 -5 4
0
0
-2 0
7 4
0
0
-2 1 -7 4
0
-2 2
2 5
0
7.49 0.81
2 -2 2
2 2 -3 5
3.03 0.22
2
2 1
2.05 0.18
2
5 5
2 2
3 5
0
-2 0
6 5
2.32 0.15
2
2
2
2 2 -4 5
0 0.2
-2
3.22 0.15
2 1 -6 5
2.51 0.22
1.89 0.17
2 2
4 5
0
-2 1
6 5
4.12 0.42
2 2 -5 5
3.64 0.29
2
7 5 16.86 0.37
2
2 3
9.06 0.56
2 3
0 3
5.34 0.42
2 2 -6 3
5.58 0.45
2 2
1 -1 3
7.72 0.35
2 0
1 4
8.73 1.58
2 3 -1 4
0
0
-2 0
10.87 0.46
2 0
2 4 10.38 0.45
2 3
0
0
-2 1 -7 5
-2 1
0 4 20.22 0.58
2 2 -6 4
7.49 0.43
2 2
2 0
1.42 1.78
2 3 -1 5
0
0
-2
0
0
-2
1 3
0
3 3
1 -2 3 1
2 3
1 -3 3 0
4 3
1
3 3
123
0
0
37.59 1.36
2 1 -1 4
4.18 0.82
4.47 0.5
2 1
1 4
6.06 0.85
32.23 1.24
2 0
3 4
0
0
0
47.85 1.57
0
-2 1 -2 4 36.00 1.4 2 1
2 4
7.45 0.76
2 0
0 4 1 5
0
2 -2
0
5 4
0
0
1 1
0 3
0
0
0
1 3
2 2 -2 2 18.26 1.69
-2 2
4.23 0.52
0
0
0
4 3
0 2
0
0
6 3
0
1 2
0
2 2
0
2 2
0
-2 1
1 -7 2
-2 2 -1 2 38.57 1.15
2.75 0.21
5 5
0
8.47 0.39
-2 2
2 5 15.25 0.57
2 3
0 5
4.76 0.28
2 2 -6 5
9.36 1.04
2
2 1 -1 5
6.66 0.24
2 0
1 6
0.29 1.85
2
2 1
7.06 0.26
2 0
2 6
1.83 0.33
2
-2 1
1 5
0 5
0
2
Cellulose (2009) 16:943–957
949
Table 3 continued h k
l
1 -4 3
rI
I
44.29 1.01
1
4 3
7.67 0.62
2 0
0 3 5 3
0 0
0 0
2 -1 3
5.11 0.31
2
0
1 3
0
2 -2 3
3.02 0.76
1 -5 3
0
2
1.65 0.87
2 3
0
m
h k
l
rI
I
2 1 -3 4 13.61 0.91 2 0
4 4
0
0
6.84 0.39 3.18 0.33
4.99 0.37
2
2 1
2 6
2.57 0.16
2
2 0
5 4
0
0
-2 1 -4 5
7.93 0.32
2 1 -3 6
2.19 0.25
2 0
4 6
0
0
3 6
4.3
0.28
2
1.88 0.16
2
2 2
1 4
0
0.2 0
0
-2 2
2 4 14.43 0.83
0
5 6
0
0
-2 2
5 6
2 3 -1 6 -2 3
0 6
2 2 -6 6
0
4 5
4.06 0.26
-2 2
0 5
0
0
-2 1
2 0
5 5
0
0
-2 1 -4 6
-2 2 -1 5
8.78 0.32
7.38 0.29
2 2
1 5
0
0
0.97 1.1
2
1.06 0.79
2
0
-2 1
0
0
-2 1 -1 8
0
0
-2 2 -1 7 2 2
1 7
2.45 0.22 0
0
1 7
0.3
1.89
2 2 -2 7
5.38 0.55
2 7
0.73 0.57
2 1 -5 7
0
2 -3 6 1 5 6
2.44 0.22 3.93 0.26
2 1 0 7 2 1 -1 7
1.05 0.34 1.14 0.29
2 2 2 7 2 2 -3 7
3.72 0.52 1.85 0.31
2
3 6
0
6 6
11.56 0.33
-2 1
1 7
1.06 0.26
2 0
3 7
0
2 -4 6
7.47 0.32
2 1 -2 7
1 -6 6
7.54 0.33
2 1
2
4 6
0
1
6 6
7.35 0.86
2 0
4 7
0
2 -5 6
8.82 0.41
2 1
3 7
1.96 0.17
0
8.97 0.38
4 7
0 8
0
2 0
2 1 -4 7
0
5 7
-2 0
-2 1
0 6
0 7
0
0
-2 2
2 2
6.32 0.44
0
2.06 0.18
-2 0
2 6
-2 1 -3 7
4 6
0
7.19 0.61
0
0
2 1
1 8
1.14 0.72
-2 0
3 8
0
2 1 -2 8 -2 1
2 8
0
2 2
2 1 -3 8 2 0 4 8
0.84 0.38 2 0 0 -2
2 1
3 8
0.89 0.52
-2 1 -4 8
1.83 0.67
2 2
5 7
2.05 0.28
3 7
0
1.97 0.43
2 0
6 7
3.28 0.28
2 1
4 8
2.16 1.19
1.75 0.2
2 2 -4 7
1.87 0.27
2 2
0 8
0
2 0
5 8
1.35 0.18 0
2 1 -6 7
2.05 0.25
-2 2
4 7
0
0
2 1
6 7
5.65 0.6
2 -2
2.79 1.28
2 1
0
2 -2
1.61 0.46
-2 2
0
2 -2
2 1
0
1 -5 6
2 7
-2 1 -2 6
2 1
2
7 6
4.41 0.19 2 0 0 -2
1 6 3 6
9.28 0.29
0
1 -7 6
2
2 1 2 0
0
3 3
0
2
3.36 0.21
3 5
2
0
1.93 0.26
4 5
9.11 0.6
0
0 6
2 1 -1 6
2 0
-2 2 -1 4 10.1
0
m
-2 1
0
0
2 1 2 5 2 1 -3 5
rI
I
0
2 2 -2 4
0
-2 1
l
3.98 0.34
2 1 -5 4
5.64 0.42
0
h k
0
0.25
0
m
0 4
0.8
1 6
0
rI
4 4
2.86 0.17
2 -2 6
3 5
I
2 1
5 3
2
2 0
l
-2 2
2 -3 3
4.42 0.19
h k
-2 1 -2 5 19.93 0.73
-2 1 3 4 8.45 0.45 -2 1 -4 4 14.23 0.52
1
2 -1 6
m
-2 2 -1 8
2
0
-2
0
0
-2
2.3
1.29
2
2
2.07 0.22
2 0
1 8
0.33 5.98
2
2.01 0.2
2 0
2 8
0.78 2.46
2
A negative multiplicity indicates that the intensity of this reflection is included in the intensity of the following reflection as an overlapping composite intensity
will always tend to slightly underestimate the background. In order to restore a flat analysis of variance and remove any systematic bias in the refinement we excluded all reflections with Fc/Fc(max) \ 0.06, where Fc is the calculated structure factor amplitude and Fc(max) is the maximum value of Fc within a particular resolution shell. This reduced the number of data from 200 to 180 and resulted in values of 0.1644 and 0.410 for R and Rx, respectively. It should be pointed out that this is similar to the conventional practice in fiber studies of leaving out very weak or absent reflections with Fo [ Fc from the
refinement in order to avoid biasing the refinement by overestimating Fo, where Fo is the observed or measured structure factor amplitude. Residual density in a Fourier difference map had a rms deviation from the mean of 0.12e/A3 and maximum and minimum peak values of 0.43 and -0.56e/A3. Using the HOPE option in SHELX to model anisotropic thermal displacement did not improve R or Rx. We did not attempt to attach H atoms to O or N atoms because of the ambiguity associated with their orientations. Repeating the initial structure refinement step with the chains in the ‘‘up’’ configuration result in values of 0.4742 and 0.8299 for R and Rx and we could not
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950
Cellulose (2009) 16:943–957 Table 4 Fractional atomic coordinates of the atoms for the asymmetric unit of EDA-cellulose I Atom
x
y
z
Uiso
Cellulose
Fig. 4 Section through Omit maps calculated using the observed X-ray amplitudes and model phases out to a ˚ , but with the hydroxymethyl group O6 resolution of 1.5 A atom omitted from the phase calculation. Blue density represents a 2Fo - Fc Omit map displayed at a threshold level of 1.3r. Red density represents a Fo - Fc Omit map displayed at a threshold level of 2.5r. The skeletal model represents the cellulose chain. One residue is shown as a ball and stick model with carbon, oxygen and hydrogen atoms explicitly represented in yellow, red and gray, respectively. Density (indicated by the blue arrow) can be clearly associated with the hydroxymethyl group in the gt position. Density (indicated by the red arrow) can also be associated with an EDA molecule
O2
0.900(11) -0.243(2)
0.057(3)
0.096(13)
O3
1.105(9)
-0.180(2)
0.319(3)
0.060(11)
O4
1.093(7)
-0.0634(18) -0.1203(18) 0.044(9)
O5
1.105(7)
0.0732(16)
0.042(2)
0.044(8)
O6
1.020(8)
0.307(2)
0.075(3)
0.041(9)
C1
0.971(7)
-0.0389(19)
C2
1.045(10) -0.1329(17)
0.096(2)
0.056(9)
C3
0.972(10) -0.1020(17)
0.232(2)
0.053(9)
C4
0.938(9)
C5
0.975(8)
0.0000(17) 0.054(10)
-0.0260(17) -0.2333(19) 0.047(8) 0.1094(17) 0.2361(15)
0.160(2) 0.183(2)
0.044(8)
C6
1.077(12)
H1
0.75658
-0.03562
-0.00710
0.06443
0.051(11)
H2
1.25845
-0.14082
0.09154
0.06730
H3 H4
0.75739 0.72467
-0.11409 -0.03569
0.24192 -0.21887
0.06330 0.05648
H5
0.75938
0.10322
0.15147
0.05286
H6A
0.97815
0.26541
0.25785
0.06064
H6B
1.28761
0.24162
0.20019
0.06064
N12
0.51(2)
0.637(4)
0.415(8)
0.19(4)
N22
0.577(19)
0.397(6)
0.547(4)
0.20(4)
C72
0.393(13)
0.520(5)
0.368(4)
0.08(2)
C82
0.594(15)
0.422(5)
0.408(4)
0.09(2)
0.52215
0.27502
0.10059
EDA
fit an EDA molecule into density in the resulting Omit map. The coordinates of the final ‘‘down’’ structure are given in Table 4 and are represented in Fig. 5. A summary of the important refinement parameters is given in Table 2. Selected conformational features of EDA-cellulose I and other cellulose allomorphs are given in Table 5. Interatomic distances of interest, including those of possible hydrogen bonds in EDA-cellulose I, are represented in Table 6.
Results and discussion The 13C CP/MAS NMR spectrum collected from Cladophora sp. cellulose before EDA treatment contains chemical shifts typical of a mixture of cellulose Ia and Ib, Fig. 2a (Imai and Sugiyama 1998). In particular, the splitting of resonance lines for carbon atoms C1, C4 and C6, indicates the presence of more than one type of glucosyl conformation, in agreement with the crystallographic structures of Ia and Ib (Nishiyama et al. 2002, 2003) and two-dimensional 13C–13C correlation NMR studies (Kono et al. 2003). The spectrum collected from EDA-cellulose I has single
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H72A 0.37608 H72B 0.19672
0.50293
0.40355
0.10059
H82A 0.53576
0.34986
0.36093
0.10447
H82B 0.79628
0.44547
0.38534
0.10447
resonance lines for each carbon atom of the cellulose backbone as well as for the guest EDA molecule at 45.6 ppm, in agreement with previous 13C CP/MAS NMR studies, Fig. 2b (Chanzy et al. 1987; Numata et al. 2003; Henrissat et al. 1987). These spectra demonstrate that the mixed cellulose Ia and Ib sample has been transformed into EDA-cellulose I. The crystallographic unit cell dimensions for EDA cellulose I reported in our preliminary analysis of the X-ray data (Wada et al. 2008), and in more detail here ˚ , b = 11.330 A ˚ , c = 10.368 A ˚ and (a = 4.546 A c = 94.017), differ from those previously reported; ˚ , b = 12.3 A ˚ , c = 10.3 A ˚ , c = 43.2) (a = 12.2 A ˚ ˚, (Creely et al. 1959) and (a = 12.87 A, b = 9.52 A ˚ c = 10.35 A, c = 118.8) (Lee et al. 1984). The unit
Cellulose (2009) 16:943–957
951 Table 6 Interatomic distances of potential hydrogen bonds of interest
Fig. 5 Section through the 2Fo - Fc map of the final EDA˚ and cellulose I structure, calculated to a resolution of 1.2 A displayed at threshold levels of 1.5r (blue) and 2.5r (red). The skeletal model represents the cellulose chain and EDA molecule. One asymmetric unit is shown as a ball and stick model with carbon, oxygen, nitrogen, and hydrogen atoms explicitly represented in yellow, red, blue and gray, respectively
Table 5 Selected conformational parameters of EDA-cellulose I compared with those of the other cellulose allomorphs and complexes v
v0
h
U
W
s
EDA-cellulose I
-92
86
117
53
Ammonia-cellulose I
-96
90
117
69 -171 15.1
173 14.5
Cellulose IIII
-92
93
116
44
Cellulose II origin Cellulose II center
-97 -94
95 87
116 115
72 -165 5.0 58 -175 10.2
Cellulose Ia upper residue
-99
94.6 116 166
-74
6.9
Cellulose Ia lower residue
-98
99.2 116 167
-75
9.4
Cellulose Ib origin
-98.5 90.5 115 170
-70 10.2
Cellulose Ib center
-88.7 94.5 116 158
-83
163 10.5
6.7
The conformation of the hydroxymethyl group is defined by two letters, the first referring to the torsion angle v (O5–C5–C6–O6) and the second to the torsion angle v0 (C4–C5–C6–O6). Thus the ideal tg and gt conformations would respectively be defined as sets of two angles (180, 60) and (60, 180). The glycosidic bond angle, s is defined by (C1–O4–C4). The glycosidic torsion angles U and W, which describe the relative orientation of adjacent glucosyl residues in the same chain are defined by (O5–C1–O1–C4) and (C1–O1–C4–C3), respectively. The sugar pucker parameter h is defined as in Cremer and Pople (1975)
N(12)
O6
3.03
N(12)
O6
2.74
N(12)
O3
3.04
N(12) N(22)
O3 O2
3.43 2.69
N(22)
O2
3.05
cell contains one cellulose chain, whereas those previously reported contain two. In the crystallographic structure there is one EDA molecule per glucosyl residue, as in the previously reported X-ray structure (Lee et al. 1984), and in agreement with our previously reported density and thermogravimetric analyses (Wada et al. 2008). The P21 space group of EDA-cellulose I has an asymmetric unit that contains only one glucosyl residue, as with cellulose IIII and ammonia cellulose I (Wada et al. 2001, 2004 and 2006). All other cellulose allomorphs that we have previously studied using X-ray and neutron crystallographic techniques, including cellulose II (Langan et al. 1999 and 2001) cellulose Ia (Nishiyama et al. 2003) and cellulose Ib (Nishiyama et al. 2002) contain two glucosyl residues in the asymmetric unit. ˚3 The unit cell volume per cellulose chain is 532.7 A 3 ˚ , 347.7 A ˚ 3, for EDA-cellulose I, whereas it is 329 A ˚ 3 for cellulose Ib, cellulose IIII and and 406.9 A ammonia-cellulose I, respectively. The crystallographic result that there is one EDA molecule per glucosyl residue disagrees with the previous 13C NMR study of Numata et al. (2003) in which only one EDA molecule per two glucosyl residues was reported. One possible explanation for this discrepancy is the exact sample preparation conditions. The EDA resonance line in Fig. 2b would appear to be very much narrower, and to have a much larger peak height relative to the other resonance lines, compared to the spectra reported by Numata et al. (2003). This may indicate a different stoichiometry and a different degree of order between the samples used in this study and those used by Numata et al. (2003) and possibly the presence of unbound EDA as previously suggested by Henrissat et al. (1987). We also investigated the possibility that a correlation between occupancy and thermal displacement parameters (Uiso) in the crystallographic refinement was affecting the refined occupancy of the EDA
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952
molecule. Although values of Uiso of the C72 and C82 atoms of EDA are similar to those of the cellulose atoms, the values for N12 and N22 are significantly larger, Table 4. When Uiso was constrained to be the same for all atoms and the structure was refined, the global value of Uiso refined to 0.051, the occupancy of the EDA molecule to 0.7, and the values of R and Rx to 0.1901 and 0.5036, respectively. However, the values of Uiso for EDA atoms will be larger than for cellulose atoms, because the molecule is smaller and less restrained. The precise value of the occupancy will therefore be higher than 0.7, in agreement with our previous density and thermogravimeteric results (Wada et al. 2008). Selected conformation parameters of EDA-cellulose I, and those of the other cellulose allomorphs and complexes whose crystallographic structures have been determined, are shown in Table 5. The relative orientation of adjacent glucosyl residues can be described by the glycosidic torsion angles U and W and the conformation of hydroxymethyl groups can be described by torsion angles v and v0 . The hydroxymethyl group of EDA-cellulose I is in the gt conformation, as it is in cellulose II, cellulose IIII, and ammonia-Cellulose I, whereas it is tg in cellulose Ia and Ib. This would appear to be confirmed by the value of 62 ppm for C6 in the 13C CP/MAS NMR spectrum collected from EDA-cellulose I, Fig. 2b, Table 1. There are significant differences between the crystallographic conformational parameters describing chains with gt hydroxymethyl groups in different cellulose allomorphs, Table 5. The conformational parameters of EDA-cellulose I are most similar to those of the center chain of cellulose II. On average, the values of /, w, v, v, h for EDA-cellulose I differ from those of the center chain of cellulose II by 4.2 whereas they differ from those of ammonia-cellulose I by 6.8, from those of cellulose IIII by 6.5 and from those of the origin chain of cellulose II by 10.9. The differences between the conformation parameters of EDA-cellulose I and those of cellulose Ia and cellulose Ib are greater. Another interesting observation is the large up field displacement ([3 ppm) of the C4 resonance line in the 13 C NMR spectrum of EDA-cellulose I from the values observed for the native cellulose allomorphs, Fig. 2 and Table 1. In EDA-cellulose I the C4 resonance line is found at 84.6 ppm, whereas in spectra recorded from
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Cellulose (2009) 16:943–957
the native cellulose allomorphs it is found in the 89– 90 ppm region, with the exception of a resonance line at 84.5 ppm that has been associated with chains at the surface of crystalline regions (Montanari et al. 2005). For other cellulose allomorphs with hydroxymethyl groups in the gt conformation, there is an up field displacement of the C4 resonance line of similar size for Na-cellulose I (85.3 ppm; Porro et al. 2007), but smaller or negligible displacements for Cellulose IIII (87.8 ppm; Table 1) and cellulose II (88.7 and 87.4 ppm; Kono et al. 2004; Dudley et al. 1983). Therefore, there would appear to be little overall correlation between a change in hydroxymethyl group conformation from tg to gt and an up field shift in the C4 resonance line. It has previously been reported that the C4 resonance line chemical shift can be correlated to the conformation of the glycosidic linkage, represented by torsion angles U and w in Table 5 (Yamamoto and Horii 1993; Atalla and VanderHart 1999). Thus it has been suggested that the presence of a double resonance line for C4 in spectra recorded from cellulose II (88.7 and 87.4 ppm) is related to a slight difference in the glycosidic conformations of the crystallographically distinct center and origin chains (Kono et al. 2004). As already noted, the conformation of EDA-cellulose I is similar to that of the center chain of cellulose II. In particular the values of U and w are similar for EDA-cellulose I and the center chain of cellulose II, whereas they are similar for cellulose I and the origin chain of cellulose II. This might indicate that, for cellulose II, the C4 resonance lines at 88.7 and 87.4 ppm can be assigned to the origin and center chains, respectively. The conformation of the glycosidic linkage in cellulose I chains and the cellulose II origin chain is relatively flat and rigid, whereas in the EDA-cellulose I chain and the cellulose II center chain it is more relaxed. Interestingly, in Na-cellulose II, where the O3-HO5 hydrogen bond that stabilizes the glycosidic linkage is broken and the linkage is allowed to relax, there is a dramatic up field shift of the C4 resonance line to 79.5 ppm (Porro et al. 2007). A relaxation of the glycosidic linkage may therefore be partly responsible for the observed EDA-cellulose I C4 resonance line shift, although other factors, such as hydrogen bonding arrangements and strengths, may also contribute. The cellulose chains in EDA-cellulose I pack in a manner that has some similarities to the packing
Cellulose (2009) 16:943–957
953
Fig. 7 A hydrophobic stack of sugar rings in EDA-cellulose I viewed down the c-axis direction and showing only N12 of EDA. Potential hydrogen bonds involving N12 and O2 and O3 of cellulose are shown as green dashed lines. C, O, H and N atoms are represented as gray, red, white and blue balls, respectively. Covalent bonds are represented as sticks
Fig. 6 Projections of the crystal structures of (top) EDAcellulose I and (bottom) ammonia-cellulose I. The chain axes point up out of the plane of display. Therefore, the positive caxis direction points out of the plane for ammonia-cellulose I (parallel up) and the negative c-axis direction points out of the plane for EDA-cellulose I (parallel down). C, O, H and N atoms are represented as gray, red, white and blue balls, respectively. Covalent bonds are represented as sticks
arrangement in ammonia-cellulose I, Fig. 6; the chains are not staggered in the c-axis direction with respect to each other and the sugar rings pack directly on top of each other in hydrophobic stacks. In ammonia-cellulose I, an ammonia molecule is in a position to form potential hydrogen bonds to O3 and O6 atoms of these stacked chains, generating an extended zigzag network of hydrogen bonds along the edges of the hydrophobic stacks, as shown in Fig. 2 of Wada et al. (2006). In EDA-cellulose I, the N12 atom would appear to be in a position to play a similar role with potential hydrogen bonds between O3 and O6 atoms of stacked chains, Fig. 7. One of ˚ the distances between N12 and O3 is too large 3.4 A but since N12 has a large thermal displacement parameter it may still represent a hydrogen bond interaction. In addition, N22 is in a position to form potential hydrogen bonds between O2 atoms of stacked chains, a hydrogen bond bridge that is not present in ammonia-cellulose I, Fig. 8. The ammonia molecule in ammonia-cellulose I, is in a position to form a bridge between sheets and between hydrophobic stacks by forming potential hydrogen bonds between O6 and O2 of chains in stacked sheets and neighboring hydrophobic stacks. In EDA-cellulose I
Fig. 8 Crystal structure of EDA-cellulose I with the c-axis titled slightly to show potential hydrogen bonds involving the EDA molecule (green dashed lines). The intrachain hydrogen bonds along the cellulose chains have been omitted for clarity. C, O, H and N atoms are represented as gray, red, white and blue balls, respectively. Covalent bonds are represented as sticks
this bridge is also present with N12 in a position to form a potential hydrogen bond to O6 and N22 in a position to form a potential hydrogen bond to O2. There are differences between the way in which chains pack in EDA-cellulose I and ammonia-cellulose I. Chains are arranged edge-to-edge in flat sheets that stack on top of each other in ammonia-cellulose I. Within the sheets there are potential hydrogen bonds between O2 and O6 atoms of neighboring chains. In EDA-cellulose I, the chains in these sheets have been stretched apart so that the potential hydrogen bonds between chains have been replaced by potential hydrogen bonds to the bridging EDA molecule; N12 has possible hydrogen bonds to O6 and N22 to O2 in neighboring chains.
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In the complexes of cellulose with EDA and ammonia, the packing interactions between cellulose chains that are facilitated by the guest molecules would appear to be similar with the exception that in EDA-cellulose I the potential intrasheet hydrogen bond has been broken and the sheets have been stretched in the b-axis direction, thus pushing the hydrophobic stacks of cellulose chains further apart. This is reflected in the fact that the unit cell ˚, dimensions of EDA-cellulose I (a = 4.546 A ˚ , c = 10.368 A ˚ and c = 94.017) and b = 11.330 A ˚ , b = 8.81 A ˚, ammonia-cellulose I (a = 4.47 A ˚ and c = 92.7) are similar, with the c = 10.34 A exception of the value of b, which is larger in EDAcellulose I. If the two C atoms of EDA are thought of as acting as spacers between the amine groups then we can speculate that a complex of cellulose with linear diamines with longer spacers such as 1,3-propylenediamine, putrescine, cadaverine, and hexamethyleneamine would also have unit cell parameters similar to those of EDA-cellulose I and ammonia-cellulose I but with the value of b correspondingly larger. As in the case of ammonia-cellulose I, with no direct information on hydrogen atom positions, it is difficult to say anything definitive about the hydrogen bonding donor/acceptor arrangement. Furthermore, the fact that the N12 and N22 atoms have relatively large atomic displacement parameters may be an indication that there is considerable conformational variation around the C72-C82 bond with the positions of the N12 and N22 atoms varying dynamically, or statically from unit cell to unit cell. The crystal structure reported here is a temporal and spatial average of crystalline regions in the sample and even if the distance between a potential hydrogen bond donor and a potential hydrogen bond acceptor is favorable for hydrogen bond formation, that is not evidence that a hydrogen bond actually exists. ˚, However, based on the O3O5 distance of 2.8 A it is highly likely that each O3 atom donates a proton in an intrachain hydrogen bond to the O5 ring atom of the next glucosyl residue. Such a bond has been observed for all other forms of cellulose we have previously examined (Nishiyama et al. 2002, 2003; Wada et al. 2004, 2006; Langan et al. 1999, 2001) and for numerous related small molecules (PeraltaInga et al. 2002). It is also likely that, because the hydroxymethyl group is in the gt conformation, this hydrogen bond is bifurcated with a major component
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Cellulose (2009) 16:943–957
to O5 and a minor component to O6 of the next glucosyl residue. This suggests that if the O3 atoms do form hydrogen bonds with EDA in EDA-cellulose I, it is likely that they accept protons in a hydrogen bond with N12, in the same way that if O3 atoms do form hydrogen bonds with ammonia in ammoniacellulose I, it is likely that they accept protons in hydrogen bonds with N. Since the hydroxyl O6 and O2 atoms cannot donate protons in hydrogen bonds to other cellulose atoms, it is also likely that if they do form hydrogen bonds, they donate protons in hydrogen bonds to N12 and N22, respectively. Thus N12 and N22 may accept protons in hydrogen bonds from the O6 and O2 atoms in one sheet and donate protons to O6 and O2 atoms in the stacked sheet, respectively. The implication is that, as in ammonia-cellulose I and cellulose IIII, there is the possibility of extended zigzag networks of hydrogen bonds that run through EDA-cellulose I. Regardless of the exact hydrogen bonding arrangement, or whether indeed EDA is involved in formal hydrogen bonds, the C72 and C82 atoms of the EDA molecule are not in equivalent crystallographic environments, which would seem to conflict with the observation of a single resonance line for the EDA molecule in the 13C CP/MAS NMR spectrum, Fig. 2b. However, when the environment of C72 and C82 are examined in detail there are some remarkable similarities. C72 and C82 from adjacent EDA molecules are the nearest non-bonded atoms to each other ˚ ), in a repulsive interaction in which their (3.7 A hydrogen atoms point almost directly toward each other, and which would be almost equivalent for either atom. The only other significant interactions that the hydrogen atoms of C72 and C82 participate in involve larger distances and are also repulsive. Furthermore, if we consider only the cellulose hydroxyl groups that are most likely to be hydrogen bonding to the EDA molecule (O2 and O6, because they can participate in no hydrogen bonds to other cellulose atoms), then the environments of N12 (attached to C72) and N22 (attached to C82) are also similar; N12 could hydrogen bond with two O6 ˚ ) with a atoms (bond lengths; 2.74 and 3.03 A O6N12O6 angle of 104.0, and N22 could hydrogen bond with two O2 atoms (bond lengths; ˚ ) with a O2N22O2 angle of 2.67 and 3.05 A 104.7. These observations may partly explain the equivalent resonance lines for C72 and C82.
Cellulose (2009) 16:943–957
However, as noted before, it may be that the N12 and N22 atoms are highly mobile, or disordered, and are not greatly constrained by hydrogen bonds, resulting in the EDA molecule being relatively unstrained by its exact crystallographic packing environment. Conversion of cellulose I to ammonia-cellulose I to cellulose IIII can be understood as ammonia molecules penetrating between hydrogen bonded sheets that are stacked through C–HO and van der Waals interactions in cellulose I. In both the Ia and Ib allomorphs of cellulose I the sheets are staggered with respect to each other in the chain axis direction. As the ammonia molecules penetrate, they lubricate the sheets causing them to slide over each other until they are no longer staggered. In addition, the hydroxymethyl groups change rotation from tg to gt. These changes allow the ammonia molecules to be accommodated at specific sites between the sheets through potential hydrogen bond interactions. The ammonia molecules form potential hydrogen bond bridges between the sheets and also along the edges of hydrophobic stacks of chains. In the case of cellulose IIII neutron crystallographic studies have allowed the presence of specific hydrogen bonds to be directly established. When the ammonia molecules are removed, making cellulose IIII, the ammonia bridge is replaced by an intersheet hydrogen bond. The hydrophobic stacks are preserved, but now the edges of chains in neighboring stacks inter-digit in a way that allows the formation of a zipper of interchain hydrogen bonds, running along the edges of these stacks. In the case of the conversion of cellulose I to EDAcellulose I, it may well be that the EDA molecules penetrate the sheets and lubricate them in the same way in the initial stages of conversion. However, at some stage the hydrogen-bonded sheets are disrupted and stretched so that the hydrophobic stacks are pushed apart. There are no direct hydrogen bonds between cellulose chains in EDA-cellulose I and the cellulose chains are in effect isolated from each other. The EDA molecules play a similar role to ammonia molecules in ammonia-cellulose I, but in addition they also adopt the role of the intrasheet hydrogen bonds between cellulose chains. One possible mechanism for the transition from the tg to the gt conformation may be a weakening of the hydrogen bonds that stabilize the hydroxymethyl group in the tg conformation. In crystalline regions of native cellulose there are intramolecular O2-HO6,
955
and intermolecular O6-HO3, hydrogen bonds that stabilize the hydroxymethyl group in the tg conformation, and that rigidify the cellulosic backbone (Nishiyama et al. 2002, 2003). However, if these hydrogen bonds are weakened by interaction with EDA, then the glycosidic linkage will be allowed to relax and the hydroxymethyl group will be free to rotate to the energetically more stable gt conformation. It would seem possible that during the formation of EDA cellulose I, EDA molecules donate protons to O3 atoms and disrupt the O6-H…O3 hydrogen bonds found in native cellulose. The hydroxymethyl groups can then rotate to the gt conformation, in which O6 can donate a proton in a hydrogen bond to EDA. Now that several crystal phases (Ia, Ib, ammoniacellulose I, EDA-cellulose I, cellulose IIII) (Nishiyama et al. 2002, 2003; Wada et al. 2001, 2004, 2006) adopted by cellulose during its solid-state conversions by EDA and ammonia have been determined, some interesting general observations can be made. In particular, we can group these crystal phases into two configurations; one in which a large number of relatively weak C–HO interactions between cellulose chains would appear to be important for stability, which we designate natural (Ia and Ib), and another in which a small number of strong hydrogen bonds between O and O atoms, or O and N atoms, would appear to be important, which we designate activated (ammonia-cellulose I, EDA-cellulose I and cellulose IIII). The introduction of amines can be thought of as a driving force for converting the natural configuration to the activated configuration and hydrothermal treatment can be thought of as a driving force for converting the activated configuration to the natural configuration. In the natural configuration the sheets of hydrogen bonded cellulose chains are staggered with respect to each other, with the hydroxymethyl groups in the tg conformation. In the activated configuration the sheets are not staggered with respect to each other and the hydroxymethyl groups are in the gt conformation. The driving forces therefore slide the sheets with respect to each other and reorient the hydroxymethyl groups in order to convert between the normal and activated configurations. Acknowledgments We thank beam line BL38B1 at the SPring-8, Japan, for use of facilities. MW was supported by a Grant-in-Aid for Scientific Research (18780131). This study was partly funded by the French Agence Nationale de la Recherche. PL was supported in part by the Office of
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956 Biological and Environmental Research of the Department of Energy, a grant from the National Institute of Medical Sciences of the National Institutes of Health (1R01GM071939-01), and a Laboratory Directed Research and Development grant from Los Alamos National Laboratory (20080001DR).
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