Oligosaccharide analogues of polysaccharides. Part 26. Mimics of cellulose I and cellulose II: Di- and monoalkynyl C-cellosides of 1,8-disubstituted anthraquinones

Full text of DOI:10.1002/hlca.200690068

Helvetica Chimica Acta – Vol. 89 (2006)
675
Oligosaccharide Analogues of Polysaccharides
Part 26
Mimics of Cellulose I and Cellulose II: Di- and Monoalkynyl C-Cellosides of
1
,8-Disubstituted Anthraquinones
by Kadiyala V. S. N. Murty, Tong Xie, Bruno Bernet, and Andrea Vasella*
Laboratorium für Organische Chemie, ETH Zürich, CH-8093 Zürich
The anthraquinone derivatives T-x-x (x=2, 4, and 8), possessing two cellobiosyl, cellotetraosyl, and
cellooctaosyl chains, respectively, C-glycosidically bonded at C(1) and C(8) were synthesised as potential
mimics of cellulose I. The anthraquinone template enforces a parallel orientation of the cellodextrin
chains at a distance corresponding to the one between the crystallographically independent chains of cel-
lulose I, and the ethynyl and buta-1,3-diynyl linker units ensure an appropriate phase shift between them.
The H-bonding of the T-x-x mimics was analysed and compared to the one of the mono-chained ana-
logues T-x and of the known cellulose II mimics N-x-x and N-x where one or two cellodextrin chains
are O-glycosidically bonded to naphthalene-1,8-diethanol, or to naphthalene-1-ethanol. The OH signals
of T-x and T-x-x in solution in (D )DMSO were assigned on the basis of DQFCOSY, HSQC, and TOCSY
6
(
only of T-4, T-4-4, and T-8-8) spectra and on a comparison with the spectra of N-x and N-x-x. Hydrogen
bonding was analysed on the basis of the chemical shift of OH groups and its temperature dependence,
1
coupling constants, SIMPLE H-NMR experiments, and ROESY spectra. T-4-4 and T-8-8 in (D )DMSO
6
appear to adopt a V-shape arrangement of the cellosyl chains, avoiding inter-chain H-bond interactions.
13
The well-resolved solid-state CP/MAS C-NMR spectra of the mono-chained T-x (x=1, 2, 4, and 8) show
13
that only T-8 is a close mimic of cellulose II. While the solid-state CP/MAS C-NMR spectrum of the C1-
symmetric diglucoside T-1-1 is well-resolved, the spectra of T-2-2 and T-4-4 show broad signals, and that
of T-8-8 is rather well resolved. The spectrum of T-8-8 resembles that of cellulose I . A comparison of the
b
X-ray powder-diffraction spectra of T-8-8 and T-8 with those of celluloses confirms that T-8-8 is a H-bond
mimic of cellulose I and T-8 one of cellulose II.
13
Surprisingly, there is little difference between the CP/MAS C-NMR spectra of the acetyl protected
mono-chained C-glycosylated anthraquinone derivatives A-x and the double-chained A-x-x (x=2, 4, and
8
). The spectra of A-4 and A-4-4 resemble strongly the one of cellulose triacetate I (CTA I). The (less
well-resolved) spectra of the cellooctaosides A-8 and A-8-8, however, resemble the one of CTA II.
13
The similarity between the solid-state CP/MAS C-NMR spectra of A-4 and A-4-4 to the one of CTA
I, and of A-8 and A-8-8 to the one of CTA II is opposite to the observations in the acetylated cellodextrin
series.
The mono-chained A-x cellulose triacetate mimics 21 (A-2), 32 (A-4), and 55 (A-8) were synthesised
by Sonogashira coupling of the cellooligosyl-ethynes 15, 28, and 50, followed by selective deacetylation.
Complete deacetylation provided the corresponding T-x mimics. The double-chained A-x-x mimics 24
(A-2-2), 35 (A-4-4), and 58 (A-8-8) were prepared from A-x by triflation and Sonogashira coupling
with the cellosyl-buta-1,3-diynes 19, 31, and 53. Their deacetylation provided the corresponding T-x-x
mimics 25, 36, and 59. The cellooligosyl-ethynes and cellooligosyl-buta-1,3-diynes required for the Sono-
gashira coupling were prepared by stepwise glycosylation of the partially O-benzylated b-cellobiosyl-
ethyne and b-cellobiosyl-buta-1,3-diyne 13 and 17, respectively, with the cellobiosyl donor 2 and the cel-
lohexaosyl donor 47.
ꢀ
2006 Verlag Helvetica Chimica Acta AG, Zürich

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Helvetica Chimica Acta – Vol. 89 (2006)
Introduction. – There are at least four polymorphs of cellulose, termed cellulose
4
I–IV [1] [2]. Common to all of them is the C conformation of the 1,4-linked b-D-glu-
1
copyranosyl moieties and the intra-chain, inter-residue O(3)ꢀH···O(5’) H-bond, as it
was reviewed on several occasions ([3][4] and refs. quoted there). Most important
are the native, metastable cellulose I and cellulose II, the most-stable polymorph [5].
They differ by their crystal packing, i.e., by the relation between adjacent chains of
the unit cell. Native cellulose I preparations of different origin show subtle differences
in their X-ray diffraction pattern that were rationalised when Atalla and VanderHart
found that samples of native cellulose are composed of two crystalline phases, cellu-
lose I and cellulose I [6]. These phases are also evidenced by solid-state CP/MAS
a
b
13
C-NMR, Raman [7], and IR spectroscopy [8], and by electron diffraction [9–11]. Cel-
luloses I and I are not only found within a single cellulose sample [11], but also along a
a
b
single microfibril [12]. Their relative amounts vary between samples of different origin.
Cellulose I -rich specimens are found in the cell walls of some algae and in bacterial
a
cellulose, and cellulose I -rich specimens in cotton, wood, and ramie fibers [11–13].
b
The currently accepted three-dimensional structure of celluloses is mainly based on
X-ray diffraction data of microcrystals and on computer modeling [14] [15]. The crystal
structure of cellulose I and I , and the H-bond patterns were extensively investigated
a
b
by Sugiyama et al. [12] and Langan and co-workers [14]. According to Sugiyama et al.,
cellulose I is a composite of a triclinic cellulose I phase with P1 symmetry and a mono-
a
A
C
H
T
R
E
U
N
G
clinic cellulose I phase with P2 symmetry [12][13][16]. According to the model of
b
1
Langan et al. [17], cellulose II (obtained from cellulose I by regeneration or mercerisa-
tion) differs from cellulose I mainly by the antiparallel orientation of the origin and
b
centre chains, the gt conformation of the glucopyranosyl units, and intersheet H-bonds.
13
The structure of celluloses was also studied by solid-state CP/MAS C-NMR spec-
13
troscopy [18]. Analysis of selectively C-labeled celluloses resulted in an unambiguous
13
assignment of the C-NMR peaks [19], while analysis of cellulose fibers from various
sources allowed to discriminate between chains in the interior and at the surface of the
crystallites. The different conformations of chains in the interior (tg conformation) and
at the surface (gt and gg conformation) reveal that the characteristic inter-residue
C(6)OH···OHC(2’) H-bonds of native celluloses are cleaved at the surface and
replaced by inter-residue C(2)OH···OHC(6’) and/or intermolecular H-bonds to H O
2
1
or to amorphous cellulose ). Chains in the interior of the crystallite of native cellulose
show a characteristic downfield shift of C(4) (88–91 ppm) and C(6) (65–67 ppm), as
compared to C(4) and C(6) of chains at the surface that resonate at 83–85 and 62.9/
61.7 ppm [20–24]. Recent calculations suggest that H-bonding leads to a shortening
of the C(4)ꢀO bonds from 1.43 to 1.36 Š and to the downfield shift of C(4) of cellu-
lose I and I [25].
a
b
Cellotetraose and cellotriose are considered valid models of cellulose II, since sin-
gle-crystal X-ray analysis of cellooligomers such as cellotetraose [26–28] and cello-
triose [29] showed that their structure resembles the one of cellulose II. There are no
similar model compounds for cellulose I. To this day, precise structural data defining
the atomic details of native cellulose are missing. For this reason, we decided to synthe-
1)
The different conformations of interior and surface chains of the crystallites were also visualised by
atomic-force microscopy [9].

Helvetica Chimica Acta – Vol. 89 (2006)
677
sise model compounds of cellulose I, opting for compounds where cellodextrin chains
of increasing length are attached in a parallel way to a suitable template.
We already described the design and synthesis of the cellooligomers N-x and N-x-x
2
(
x=2, 3, 4, and 8) ) where the cellooligosyl chains are glycosidically attached to naph-
thalene-1-ethanol and naphthalene-1,8-diethanol (cf. Fig. 2) [30]. However, solid-state
13
CP/MAS C-NMR spectroscopy characterised the bis-cellooctaoside N-8-8 as a mimic
of cellulose II rather than of cellulose I, showing that a parallel attachment of two cel-
looligosaccharide chains to naphthalene-1,8-diethanol is not sufficient to generate a
model for cellulose I or I [4]. It appeared necessary to design a templated model
a
b
that assures a parallel orientation of the chains, and that mimicks not only the distance
between the origin and centre chains of cellulose I but also the phase shift between
them. MM3* Calculations (Macromodel V. 6, gas phase [31]) showed that 1-(buta-
1
,3-diynyl)-8-ethynylanthraquinone fulfils these conditions. It fixes the chains in a par-
allel orientation, and leads to a distance between two C-glycosidically attached cello-
oligosaccharide chains of 6.02 Š and to an appropriate phase shift of 2.59 Š, as dis-
ACHTREUNG
cussed before [3]. A further advantage is the anticipated difference of the chemical
shift of the NMR signals of the template and of the cellooligosaccharide chains.
We planned to investigate the mono- and the double-chained cellobiosyl, cellote-
traosyl, and cellooctaosyl anthraquinonyl derivatives; this should allow to also study
the influence of the chain length on the intramolecular, inter-chain interactions. The
synthesis of the symmetric and asymmetric b-D-glucopyranosyl-ethynylated and
-
buta-1,3-diynylated anthraquinones, and the analysis of their H-bonding in solution
were published [3]. We now describe the synthesis of the corresponding templated
mono- and double-chained cellooligosides and the investigation of their H-bonding
in the solid state and in DMSO solution; we also describe the synthesis and spectra
of the corresponding analogues of cellulose triacetate (CTA).
Results and Discussion. – 1. Synthesis of b-Cellobiosyl-ethyne and b-Cellobiosyl-
buta-1,3-diynes. We planned to synthesise the templated even-numbered cellooligo-
sides (exemplified by the cellobiose derivative A) by coupling the corresponding eth-
ynyl and butadiynyl C-glycosides, such as the protected cellobiosyl-ethyne and cellobio-
syl-buta-1,3-diyne units C and D to an anthraquinone moiety B, as described for the
synthesis of the corresponding glucosyl analogue [3] (Fig. 1). The required cellobiosyl
C-glycosides should be readily available via a protected cellobionolactone E that
should be obtained from commercial cellobiose octaacetate 1 [30].
3
Cellobiose octaacetate (1) was treated with HBr in AcOH to give 94% ) of the
known bromide 2 [32] (Scheme 1). Glycosidation of allyl alcohol with 2 in the presence
of Hg(CN) yielded 85% of the acetylated b-cellobioside 3 [33]. Following the proce-
2
dure of Vliegenthart and co-workers [34], 3 was deacetylated (MeONa in MeOH;
yield >98%). The resulting allyl cellobioside 4 was transformed into the benzylidene
acetal 5 (89%) [35][36] and further in 73% yield to the benzylated 6 [34][36]. To obtain
higher melting derivatives we also transformed 5 to the 4-chlorobenzyl ether 7 (83%)
with a melting point ca. 208 higher than that of 6. Regioselective reductive cleavage of
2
)
)
For the numbering of these cellosides, see Results and Discussion, Chapt. 4.
The yield of 2 was 85–80% when the reaction was performed on a scale >8 0 g.
3

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Helvetica Chimica Acta – Vol. 89 (2006)
Fig. 1. Retrosynthesis of the template-bound cellobiosyl-acetylenes and cellobiosyl-buta-1,3-diynes
the 1,3-dioxane ring of the benzylidene acetal 7 with NaBH CN and HCl [30] gave a
3
mixture that was separated by flash chromatography (FC) to provide 69% of the
desired secondary alcohol 8. Reduction of 7 with BH ·Me
A
T
E
N
A
T
E
N
3
2
3
by FC gave 8 in a lower yield (60%), while reduction of 7 with Et
A
T
E
N
A
T
E
N
3
3
[
38] yielded 71% of 8. Methoxymethylation of 8 gave 9 (70%) besides 17% of starting
material. However, this derivative could not be converted satisfactorily to the corre-
sponding lactone. Deallylation of 9 under standard conditions [34] followed by treat-
ment with DMSO and Ac O [39] or with Dess–Martin periodinane [40] gave complex
2
mixtures. We, therefore, isomerised the benzylidenated allyl glycoside 7 according to
Baudry et al. [41], and cleaved the resulting prop-1-enyl glycosides with I in THF/
2
H O to obtain 87% of the hemiacetals 10. Oxidation of 10 with DMSO and Ac O
2
2
[
39] provided 97% of crude lactone 11.
Stereoselective methods for the preparation of b-D-hexopyranosyl-alkynes from
hexonolactones are well-documented [42–46]. Treatment of the cellobionolactone 11
with 1 equiv. of lithium (trimethylsilyl)acetylide (LiCꢁCSiMe ) in THF led to only lit-
3
tle conversion (Scheme 2). Slightly better results were obtained by using up to 2 equiv.
of LiCꢁCSiMe , in agreement with the reported sluggish addition of alkynyl metal
3
reagents to cellobionolactones [47]. Cerium [45][48] or aluminium acetylides [49] per-
formed hardly better, even when 2 equiv. of the reagent were used, and the same result
was obtained upon adding LiCꢁCSiMe in the presence of Lewis acids, such as scan-
3
dium or ytterbium triflate, in either THF or Et O. However, treatment of 11 with 4.5
A
H
R
U
G
2
equiv. of LiCꢁCSiMe led to complete conversion to 12 (a/b 55 :45). Performing this
3
addition on a multigram scale followed by reductive dehydroxylation and concomitant

Helvetica Chimica Acta – Vol. 89 (2006)
679
Scheme 1
a) HBr in AcOH, CHCl ; 94%. b) Hg(CN) , allyl alcohol; 85%. c) MeONa, MeOH; >98%. d)
3
2
PhCH(OMe) , MeCN, TsOH·H O; 89%. e) BnBr, NaH, DMF; 73%. f) 4-ClC H CH Cl, NaH,
2
2
6
4
2
DMF; 83%. g) NaBH CN, HCl in Et
A
H
R
U
G
A
H
R
N
A
H
R
U
G
3
2
2
(
C H )]PF , H , THF, then I in H O; 87%. j) Ac O, DMSO; 97% of crude 11.
8 12 6 2 2 2 2
reductive opening of the 1,3-dioxane ring with Et3
of the b-configured alcohol 13. C-Desilylation of 13 to 14 (MeONa in MeOH [50],
9%), followed by acetolytic debenzylation (TMSOTf in Ac O) gave the crystalline
A
H
R
U
G
A
T
E
N
3
8
2
cellobiosyl-acetylene 15 (76%).
The butadiyne 19 was prepared in a similar way. Addition of the lithium acetylide
derived from bis[1,4-(trimethylsilyl)buta-1,3-diyne] to the cellobionolactone 11 gave
the hemiketals 16 (a/b 3 :2). Concomitant dehydroxylation of 16 and reductive dioxane
ring opening yielded 52% of the cellobiosyl-buta-1,3-diyne 17. In contradistinction to
the preparation of the alkyne 12, the preparation of the butadiyne 16 required only a
slight excess of the alkynyl reagent. C-Desilylation of 17 to 18 (87%) followed by ace-
tolytic debenzylation gave the crystalline cellobiosyl-buta-1,3-diyne 19 (63%).
Sonogashira coupling [51–53] of the anthraquinone triflate 20 [3] and the cellobio-
sylacetylene 15 at 608 followed by selective deacetylation of the phenolic AcO group
(
(NH ) CO in DMF [3][54][55]) gave 74% of the C-glycoside 21 that was triflated
4 2 3
under standard conditions to yield 96% of the aryl triflate 22 (Scheme 3). Sonogashira
coupling of 22 with the cellobiosyl butadiyne 19 at room temperature [52][53] yielded
87% of the bis-C-cellobiosylated anthraquinone 24. Deacetylation of 21 with MeONa
in MeOH gave 95% of the mono-C-glycoside 23. The corresponding bis-C-cellobiosyl
derivative 25 was obtained in a yield of 71% by treating 24 with KCN in MeOH/CH Cl
2
2
[
3] followed by reversed-phase column chromatography.
1
13
The H- and C-NMR data for the glucosyl units of the cellobiosyl-ethynes 13–15,
the cellobiosyl-buta-1,3-diynes 17–19, and their template-bound derivatives 21–25 are

6
80
Helvetica Chimica Acta – Vol. 89 (2006)
Scheme 2
a) BuLi, Me3
A
H
R
U
G
A
H
R
U
G
A
H
R
U
3
3
BF ·OEt , CH Cl /MeCN; 72% of 13 from 11; 52% of 17 from 11. d) MeONa, THF/MeOH; 89% of
3
2
2
2
1
4; 87% of 18. e) Ac O, Me SiOTf, 20–258; 76% of 15; 63% of 19.
A
H
R
U
G
2
3
3
5
listed in Tables 5–8 in the Exper. Part. J(1,ꢁCH) of the ethyne 14 is larger than J(1,ꢁ
CH) of the buta-1,3-diyne 18 (2.2 vs. 0.6 Hz). J(1,2), J(2,3), J(3,4), and J(4,5) values evi-
dence the C conformation of all pyranosyl units. The ethynyl and the buta-1,3-diynyl
group have a similar influence upon the H- and C-NMR chemical shifts of the sugar
4
1
1
13
residues, as evidenced by the chemical shift differences (Dd values) for the correpond-
I
I
ing pairs 13/17, 14/18, and 15/19 (HꢀC(1 ): 0.03–0.06, other HꢀC: ꢂ0.03, C(1 ):
I
4
0
.1–0.4, C(2 ): 0.5–0.7, and other C: ꢂ0.2 ppm) ). However, the anthraquinonyl moi-
1
ety of 21 leads to a significant downfield shift, as evidenced by the H-NMR Dd values
I
I
for the pair 21 and 16. Large downfield shifts are observed for HꢀC(1 ) and HꢀC(2 )
I
I
(
0.42 and 0.25 ppm, resp.) and smaller ones for HꢀC(3 ) to HꢀC(6 ) (0.08–0.12 ppm).
II II
Even HꢀC(1 ) to HꢀC(6 ) are shifted downfield by 0.04–0.06 ppm. Similarly, the C-
I
I
I
I
signals of 21 are shifted downfield (Dd for C(1 ): 1.1, C(2 ): 0.2, C(3 ): 0.6, and C(4 ) to
II
I
C(6 ): ꢂ0.1 ppm). C(1 ) of the cellobiosyl-ethynes and cellobiosyl-buta-1,3-diynes res-
4)
According to the 1996 recommendations of the nomenclature of carbohydrates, the monomer units
of the cellooligosides are numbered with roman numerals in ascending order starting from the alky-
nylated end (corresponding to the reducing end).

Helvetica Chimica Acta – Vol. 89 (2006)
681
Scheme 3
a) [Pd(PPh ) ]Cl , CuI, Et N/DMF 1:5, 608, then (NH ) CO ; 74%. b) As a, but at 248 and without
A
H
R
U
G
3
2
2
3
4
2
3
(
NH ) CO ; 87%. c) Tf
A
H
R
U
G
A
T
E
N
4
2
3
2
7
0%.
I
onates upfield at 68.3–70.4 ppm, whereas the signal for C(5 ) is shifted downfield to
7
6.8–79.3 ppm (compare with 74.9 ppm for the b-celloside 7). The position of the
C-NMR signals of the ethynyl and the buta-1,3-diynyl C-atoms is characteristically
1
3
influenced by the terminal substituent (Me Si, H, or anthraquinonyl). The assignment
A
H
R
U
G
3
1
13
of the H- and C-NMR signals of 24 and 25 to the ethynylated or buta-1,3-diynylated
chain is based on the interpretation of DQFCOSY and HSQC spectra and on a com-
parison with the values of the mono-chained cellobiosyl-ethynes 21 and 23, and the
mono- and double-chained glucopyranosyl-ethynes and glucopyranosyl-buta-1,3-
diynes (data in [3]). Due to the shorter distance to the anthraquinonyl moiety, the
1
I
H-NMR signals of the ethynylated chain of 24 (except the one for HꢀC(4 )) are shifted
downfield relative to the corresponding signals of the buta-1,3-diynylated chain; stron-
I
I
I
gest downfield shifts are observed for HꢀC(1 ), HꢀC(2 ), and HꢀC(5 ) (Dd=0.25,
.18, and 0.10 ppm, respectively).
. Synthesis of the Template-Bound Cellotetraosyl-ethynes and Cellotetraosyl-1,3-
0
2
butadiynes. We planned to prepare the intermediate acetylated cellotetraose C-glyco-
sides 28 and 31 via 26 and 29 that should be obtained by Koenigs–Knorr glycosylation
[
56] with the cellobiosyl bromide 2 of the selectively protected cellobiosyl-ethyne 13
and cellobiosyl-buta-1,3-diyne 17, respectively (Scheme 4). Glycosidation in toluene/
CH Cl 1:1 with 4 equiv. of 2 at ꢀ358 to 258 for 2 days and in the presence of excess
2
2
AgOTf yielded 84% of the crystalline cellotetraosyl-ethyne 26, while the AgOTf-pro-
moted glycosidation of 13 by 2 in ClCH
A
T
E
N
CH
A
H
E
N
Cl at ꢀ358 gave only traces of 26, and
2
2
changing the solvent to CH Cl or MeCN hardly improved the conversion. C-Desilyla-
2
2

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82
Helvetica Chimica Acta – Vol. 89 (2006)
Scheme 4
a) AgOTf, 3-Š mol. sieves, toluene/CH Cl ; 84% of 26; 82% of 29. b) Bu NF·3 H O, THF; 86% of
A
H
R
U
G
2
2
4 2
2
7; 94% of 30. c) Me3SiOTf, Ac O; 76% of 28; 65% of 31.
A
H
R
U
G
2
tion of 26 with Bu NF·3 H O afforded 86% of 27. Acetolytic debenzylation of 27 gave
A
H
E
N
4 2
the crystalline peracetylated cellotetraosyl-ethyne 28 (76%).
The cellotetraosyl-buta-1,3-diyne 31 was similarly synthesised from the bromide 2
and the cellobiosyl-buta-1,3-diyne 17. Glycosylation of 17 with 2 under the same con-
ditions that we used for the synthesis of 26 yielded 82% of 29. Desilylation of 29 gave
the terminal butadiyne 30 (94%) that was transformed by acetolysis to the peracetate
3
1 (65%).
Sonogashira coupling of the triflated anthraquinone 20 with the cellotetraosyl-acet-
ylene 28 under the conditions used for the preparation of the cellobiosyl-ethyne 21 and

Helvetica Chimica Acta – Vol. 89 (2006)
683
the cellobiosyl-buta-1,3-diyne 24 produced a complex mixture (Scheme 5). Only little
conversion resulted from performing the coupling at ambient temperature. Addition
of Bu4
A
H
R
U
G
[
57], and the coupling of 20 with the cellotetraosyl-acetylene 28 occurred cleanly at
5
ambient temperature ). Selective deacetylation of the crude coupling product and
FC gave 75% of the cystalline cellotetraosylated 1-hydroxyanthraquinone 32. Triflation
under standard conditions transformed 32 to 33 (95%). The Bu NI-promoted coupling
A
H
R
U
4
of this triflate with the cellotetraosyl-1,3-butadiyne 31 gave the crystalline bis-cellote-
traoside 35 (59%). The mono-cellotetraoside 32 was deacetylated with MeONa in
MeOH to provide 95% of 34, while complete deacetylation of the bis-cellotetraoside
35 to 36 proved difficult. Standard methods, such as treatment with MeONa in
MeOH, K CO in MeOH, or KCN in MeOH failed to provide the desired product.
2
3
Deacetylation with aqueous solutions of trialkylamines, such as Et N led to a very
A
H
R
U
G
3
poor conversion even after 7 days, while aqueous KOH (30 equiv.) provided only
small amounts of 36. Finally, 78% of the completely deacetylated 36 were obtained
by using 28–29 equiv. of Bu NOH in H O.
A
H
R
U
G
4 2
1
13
The H- and C-NMR data for the glucosyl units of the cellotetraosyl-ethynes and
-
buta-1,3-diynes 26–36 are listed in Tables 9–12 in the Exper. Part. The acetoxylated
and benzyloxylated centres of 26, 27, 29, and 30 show the expected relative chemical
1
13
shifts ( H-NMR: downfield shift for acetoxylated centres, C-NMR: downfield shift
1
13
for benzyloxylated centres). The H- and C-NMR chemical shifts for unit I and IV
of the cellotetraosyl-ethynes and -buta-1,3-diynes 28 and 31–36 agree well with the cor-
responding values for unit I and II of the peracetylated cellobiosyl-ethynes and -buta-
1
13
1
,3-diynes. Unit II and III of 28 and 31–33 show slightly shifted H- and C-NMR sig-
1
nals ( H-NMR: Ddꢂ0.03 ppm with the exception of 0.06–0.08ppm for H ꢀC(2) of 28
13
1
and 31–33 and for HꢀC(5) of 32 and 33; C-NMR: Dd<0.2 ppm). Individual H- and
13
C-NMR signals are assigned to unit I of the ethynylated and buta-1,3-diynylated chain
of 35, whereas the signals for units II to IV of these chains overlap increasingly with the
distance of the centres from the anthraquinonyl moiety.
3
. Synthesis of the Template-Bound Cellooctaosyl-ethynes and Cellooctaosyl-buta-
1,3-diynes. The preparation of the cellooctaosyl-alkynes according to the route descri-
bed above for 36 requires a cellohexaosyl donor. Such a donor should be accessible in
sufficiently large amounts by two sequential glycosidations, starting with cellobiosyl
IV
derivatives. The synthesis of the HOꢀC(4 ) unprotected cellotetraoside 43 from the
cellobiosyl bromide 2 and the cellobiosyl acceptor 38 [34][36] was described [30]
(Scheme 6). The key step of this synthesis – the AgOTf-promoted Koenigs–Knorr gly-
cosidation of 38 with 1.2 equiv. of 2 in ClCH CH Cl at ꢀ308 – afforded 89% of the cel-
2
2
lotetraoside 39. We could not reproduce the high yield by following this procedure.
Even using 3 equiv. of 2 and changing the solvent to toluene/CH Cl yielded only
2
2
50% of 39. However, 39 was obtained in 87% yield by a TMSOTf-promoted glycosida-
tion of 36 with the trichloroacetimidate 37 [61] in CH Cl at ꢀ408. The next steps, i.e.,
2
2
deacetylation to 40, benzylidenation to 41, benzylation to 42, and regioselective reduc-
tion to the alcohol 43 were performed according to [30], and resulted in an overall yield
5)
For other Bu NI-promoted cross couplings, see [58][59] and refs. cited therein. For a CuI-promoted
A
H
R
U
G
4
Sonogashira coupling, see [53][60].

6
84
Helvetica Chimica Acta – Vol. 89 (2006)
Scheme 5
a) [Pd(PPh ) ]Cl , CuI, Bu
A
H
R
U
G
A
H
R
U
G
3
2
2
4
(
NH ) CO ; 59%. c) Tf
A
H
R
U
O, Et
A
H
R
U
A
H
R
U
4
2
3
2
3
2
2
of 71%. The TMSOTf-promoted glycosidation of 43 with the trichloroacetamidate 37
in CH Cl at ꢀ608 gave the cellohexaoside 44 (94%). Deallylation of 44 followed by
2
2
6
hydrogenolytic debenzylation and acetylation yielded 75% of a/b-45 2 :3 ). Selective
deacetylation of the anomeric AcO group of a/b-45 with (NH ) CO in DMF [54]
4
2
3
gave the hemiacetals a/b-46 2 :1 (78%) [66] that were treated with Cl CCN in the pres-
3
ence of DBU. Crystallisation of the product from AcOEt/hexane yielded 71% of a 94 :6
mixture of the trichloroacetimidates a/b-47 [66]. The cellooctaoside 48 was obtained in
8
8% by a BF ·OEt -promoted glycosylation of a slight excess of the cellobiosyl-ethyne
3 2
13 with a/b-47 94 :6, while the TMSOTf-promoted glycosylation of 13 with a slight
excess of a/b-47 94 :6 at ꢀ788 yielded only 40% of 48. C-Desilylation of 48 with
Bu NF·3 H O to 49 followed by acetolytic debenzylation provided the peracetylated
AHCTREUNG
4
2
cellooctaosyl-ethyne 50 (75%). Similarly, the cellooctaosyl-buta-1,3-diyne 53 was syn-
thesised via 51 and 52 from a/b-47 94 :6 and the cellobiosyl-buta-1,3-diyne 17 (42%
overall yield).
The Bu NI-promoted Sonogashira coupling at 0–288 of the triflate 20 with the cel-
A
H
R
U
G
4
looctaosyl-ethyne 50 gave 83% of the octaosylated anthraquinone 54 (Scheme 7).
Selective deacetylation of the aromatic AcO group of 54 gave the hydroxy-anthraqui-
none 55 (88%) which was triflated to 56 (59%). The Bu NI-promoted coupling of 56
A
H
R
U
G
4
with the cellooctaosyl-1,3-butadiyne 53 gave 40% of the bis-cellooctaoside 58. The
6)
For the synthesis and NMR data of a-45, see [62–65].

Helvetica Chimica Acta – Vol. 89 (2006)
685
Scheme 6
a) Me3
A
H
R
U
G
2
2
MeOH; 99%. c) ZnCl , PhCHO; 84%. d) BnBr, NaH, DMF; 93%. e) NaBH CN, HCl, 3-Š mol.
2
3
sieves, THF; 92%. f) H , [Ir(MePh
A
H
R
U
G
A
T
E
N
2
2
AcOEt/MeOH; Ac O, pyridine; 75% of a/b-45 2 :3. g) (NH ) CO , DMF; 78% of a/b-46 2 :1. h)
2
4
2
3
Cl CCN, DBU, CH Cl ; 71% of a/b-47 94 :6. i) BF ·Et
A
H
R
U
G
3
2
2
3
2
2
2
4
8; 72% of 51. j) Bu4NF·3 H O, THF; 92% of 49; 83% of 52. k) BF ·OEt , Ac O; 82 of 50; 70% of
A
H
R
U
G
2
3
2
2
5
3.

6
86
Helvetica Chimica Acta – Vol. 89 (2006)
Scheme 7
a) Pd(PPh ) Cl , CuI, Bu
A
H
R
U
G
A
H
R
U
G
3
2
2
4
c) Tf2
A
H
R
U
G
A
H
R
N
A
H
R
U
G
3
2
2
mono- and the bis-octaosides 55 and 58 were deacetylated with aqueous Bu NOH,
A
H
E
N
4
similary to 35, but sonification was required to dissolve the octaosides. This provided
the deprotected mono-cellooctaoside 57 in 43% and the deprotected bis-cellooctaoside
5
9 in 55% yield.
1
13
The H- and C-NMR chemical shifts for unit I and VIII of the cellooctaosyl-
ethynes and cellooctaosyl-buta-1,3-diynes 50, 53–55, and 57–59 resemble those of
the terminal units of the corresponding cellobiosyl and cellotetraosyl analogues,
1
13
whereas the H- and C-NMR signals for units II to VII of the cellooctaosyl derivatives
mostly overlap (see Tables 13 and 14 in the Exper. Part).
4
. Are the Template-Bound Cellooligosyl C-Glycosides Mimics of Cellulose I? Com-
parative Analysis of the Template-Bound Cellooligosyl-ethynes and Cellooligosyl-buta-
,3-diynes. To facilitate the discussion, we have replaced the compound numbers of the
1
oligosides of interest, designating the mono- and double-chained anthraquinonyl deriv-
atives as T-x and T-x-x with x and x-x (x=2, 4, and 8) denoting the number of glucosyl
residues (Fig. 2,a). Similarly, the peracetates are labeled A-x and A-x-x. The depro-
tected 2-naphthylethyl cellooligosides [4], of interest as reference compounds, are de-
A
C
H
T
R
E
U
N
G
signated as N-x and N-x-x. The ethynylated chain of A-x-x and T-x-x is termed E and
the buta-1,3-diynylated chain B. According to carbohydrate nomenclature, the gluco-
pyranosyl units of the cellooligosyl chains are labeled with roman numerals, as
4
shown in Fig. 2,b, for the cellotetraoside T-4-4 ). However, as the NMR signals of
the internal units overlap strongly, particularly in the octaoside derivatives, we desig-

Helvetica Chimica Acta – Vol. 89 (2006)
687
Fig. 2. a) Numbering of the template-bound single- and double-chained cellodextrins, and labeling of
the chains of T-x-x, and b) labeling of the glucopyranosyl units of template-bound cellodextrins
nate units I as a, the terminal units as c, and the sum of the internal units (units II and III
of tetraosides and units II–VII of octaosides) as b. These designations of the glucosyl
residues reflect regularities in the NMR spectra, as discussed below. To further simplify
the notation, the O-atoms are numbered in the same way as C-atoms.
The template-bound cellooligosaccharides T-x and T-x-x (x=2, 4, and 8) are yellow-
to-brownish solids that decompose upon melting. The double-chained T-x-x possess
higher melting points than their mono-chained T-x analogues. The shorter T-x and
T-x-x (x=2 and 4) are well soluble in DMSO, DMF, and N,N-dimethylacetamide
(
>40 mmol/l), slightly soluble in H O (<5 mmol/l), and nearly insoluble in MeOH,
2
EtOH, CF CH OH, AcOH, THF, dioxane, and CHCl . The solubility of T-x and T-x-x
3
2
3
in H O decreases with increasing number of sugar residues. The cellooctaosides T-8
2
and T-8-8 are only soluble in DMSO. Hence, most of the NMR studies were conducted
in (D )DMSO solution.
6

6
88
Helvetica Chimica Acta – Vol. 89 (2006)
1
4
.1. Investigation of the H-Bonding of T-x and T-x-x in DMSO Solution. The H-
NMR chemical shift of the OH groups (d(OH)), the vicinal coupling constant
J(H,OH)), and the temperature dependence of the OH signals (Dd(OH)/DT) are use-
ful parameters for the investigation of H-bonding of alcohols and polyols in (D )DMSO
(
6
[
(
67][68]. Fully solvated OH groups acting as H-donors in an intermolecular H-bond to
D )DMSO are characterised by a downfield shift of the OH signal, J(H,OH) values of
6
4.5–5.5 Hz for equatorial and of 4.2–4.4 Hz for axial OH groups, and a strong temper-
ature dependence (jDd(OH)/DTj>4.5 ppb/K). OH Groups acting as H-donors in an
intramolecular H-bond are readily detected by an upfield shift of the OH signals, by
J(H,OH) values deviating from those of fully solvated OH groups, and by a weak tem-
perature dependence (jDd(OH)/DTj<3 ppb/K) [67][68]. The d(OH) values for OH
groups of monosaccharides and of the terminal units of oligosaccharides are a useful
reference for the interpretation of d(OH) values for OH groups of the internal units
of oligosaccharides. The H-bonding of b-cellobiose and methyl b-cellobioside [69] in
I
(
D )DMSO has been analysed [68]. All OH groups, with the exception of HO(3 )
6
are more or less fully solvated. A completely persistent inter-residue O(3)ꢀH···O(5’)
H-bond of methyl b-cellobioside is evidenced by J(3,OH)=1.7 Hz, d(HO(3))=4.68
ppm, and Dd(HO(3))/DT=ꢀ2.6 ppb/K.
In the templated cellooligosaccharides N-x and N-x-x (x=2, 3, 4, and 8) one or two
glycosidically bonded cellodextrin chains are attached to naphthalene-1-ethanol and
naphthalene-1,8-diethanol. The H-bonding of solutions of these oligosaccharides in
(
D )DMSO was analysed [4]. HO(3a) and HO(3b) (d(OH)=4.59–4.73 ppm,
6
J(H,OH)<2 Hz, Dd(OH)/DT=ꢀ1.7 to ꢀ2.2 ppb/K) are engaged in inter-residue H-
bonds to O(5’) of the neighbouring glucosyl unit, whereas the other OH groups are
involved in intermolecular H-bonds to the solvent (J(2,OH)=J(3c,OH)=
J(4c,OH)=4.5–5.3 Hz, J(6,OH)=5.4–6.5 Hz, Dd(OH)/DT=ꢀ4.2 to ꢀ7.2 ppb/K).
HO(2b) resonates at lowest field (5.37 ppm), followed by HO(2c) (5.20 ppm),
HO(2a) (5.13–5.14 ppm), HO(3c) (4.99 ppm), and HO(4c) (4.96 ppm), whereas
HO(6b) resonates at 4.63–4.69 and HO(6a) and HO(6c) at 4.52–4.61 ppm. Only
small shift differences (Dd(OH) values) were observed for the OH groups of the inter-
III
nal units b (Dd(OH)ꢂ0.01 ppm with the exception of 0.08ppm for HO(3 ) of the cel-
VII
lotetraosides and HO(3 ) of the cellooctaosides). Weak interchain H-bond interac-
tions in N-x-x were only observed for unit a closest to the template.
No concentration dependence of the d(OH) and d(HꢀC(1)) values of T-4-4 was
observed in (D )DMSO solution at concentrations between 18and 62 mmol ( Fig. 3).
6
1
Since all H-NMR spectra of T-x and T-x-x were recorded at low concentration (10
mmol/l for T-x and 5 mmol/l for T-x-x), solute–solute interactions can be neglected.
All OH groups of the monoglucoside T-1 and the diglucoside T-1-1 are solvated in
(
D )DMSO [3]. A weakly persistent flip-flop H-bond between the two primary OH
6
groups of the diglucoside T-1-1 is suggested by the upfield shift of both OH groups
(0.04 ppm for HO(6E) and 0.09 ppm for HO(6B) relative to HO(6) of T-1).
The unambiguous assignment of the OH signals of T-x and T-x-x (x=2, 4, and 8) in
(
D )DMSO is based on the interpretation of DQFCOSY, HSQC, and TOCSY (only of
6
T-4, T-4-4, and T-8-8) spectra and a comparison with the spectra of N-x and N-x-x
Table 1). The OH groups of T-x and T-x-x show similar chemical shifts as the corre-
sponding OH groups of N-x and N-x-x except for HO(2aE), HO(2aB), and HO(6a)
(

Helvetica Chimica Acta – Vol. 89 (2006)
689
1
Fig. 3. Concentration-dependence of the H-NMR spectra of T-4-4 in (D )DMSO: HꢀC(1) and OH
6
signals
which are shifted downfield by 0.43, 0.54, and 0.10 ppm, respectively, due to the alkynyl
substituent [70]. HO(3a) and HO(3b) of T-x and T-x-x are involved in an inter-residue
H-bond to O(5’) as evidenced by the upfield shift (4.75–4.65 ppm) and a small J(3,OH)
value (<1.5 Hz). The other OH groups of T-x and T-x-x are engaged in intermolecular
III
H-bonds to (D )DMSO. As it was already observed for N-4 and N-4-4, HO(3 ) of T-4
6
II
and T-4-4 resonates downfield to HO(3 ) (Dd=0.06–0.07 ppm). A similar downfield
shift is expected for HO(3 ) of T-8 and T-8-8, but overlapping HO(3) and HO(6) sig-
VII
nals prevent an exact assignment.
A comparison of d(OH) of the ethynylated chain E of the di-cellosides T-x-x (x=2,
4
, and 8) with d(OH) of T-x (x=2, 4, and 8; Ddꢂ0.04 ppm; Table 1) shows a weak influ-
ence of the buta-1,3-diynylated chain of T-x-x on the chemical shift of the OH groups of
the E chain. These small shift differences suggest at best very weak inter-chain H-bond
interactions. The OH groups of the buta-1,3-diynylated chain B of T-x-x (x=2, 4, and 8)
possess similar chemical shifts as the OH groups of the ethynylated chain E (Ddꢂ0.03
ppm) with the exception of HO(2a) which shows the expected downfield shift due to
the buta-1,3-diynyl substituent (0.21 vs. predicted 0.16 ppm [70]). Duplication of OH
signals of T-x-x is only observed for those OH groups that are close to the template,
II
i.e., OH groups of unit a and HOꢀC(2 ) (part of units b). This restricts the analysis
1
by H-NMR spectroscopy to inter-chain H-bond interactions of these OH groups of
T-x-x.
HO(2a) of T-x (x=2, 4, and 8) resonates as broad singlet at 5.56–5.58ppm, whereas
HO(2aE) and HO(2aB) of T-x-x give rise to sharp doublets at 5.56–5.57 and 5.77–5.78
ppm, respectively (Table 1). That HO(2a) of T-x and HO(2aE) of T-x-x display the

6
90
Helvetica Chimica Acta – Vol. 89 (2006)
1
Table 1. H-NMR d(OH) [ppm] and J(H,OH) [Hz] Values (in parentheses) of T-x and T-x-x (x=2, 4, 8) in
(
D )DMSO (assignments based on DQFCOSY and HSQC spectra and on TOCSY spectra of T-4, T-4-4,
6
and T-8-8)
T-2
T-4
T-8
E chain
E chain
E chain
HO(2a) 5.58a)
HO(3a) 4.82 (<1.5)
HO(6a) 4.71 (5.8)
HO(2b)
HO(3b)
HO(6b)
5.58a)
4.76 (<1.5)
4.68 (5.1)
5.42 (5.0), 5.43 (5.0)
4.75 (<1.5), 4.69 (<1.5)
4.71 (5.2)
5.56a)
4.74–4.66
4.74–4.66
5.41 (4.9), 5.39 (ca. 4.4; 6 H)
4.74–4.66
4.74–4.66
HO(2c) 5.27 (4.7)
HO(3c) 5.05 (4.6)
HO(4c) 5.02 (4.7)
HO(6c) 4.64 (5.2)
5.24 (4.7)
5.04 (3.8)
5.00 (4.9)
4.61 (5.1)
5.21 (4.8)
5.01 (4.8)
4.98(5.3)
4.58(5.1)
T-2-2
T-4-4
T-8-8
E chain, B chain
E chain, B chain
E chain, B chain
HO(2a) 5.57 (5.5), 5.78 (5.8)
HO(3a) 4.80 (<1.5), 4.81 (<1.5) 4.74 (<1.5)
5.56 (5.7), 5.77 (5.9)
5.56 (5.5), 5.77 (5.8)
4.74–4.65
HO(6a) 4.71 (6.0)
HO(2b)
4.72–4.65
4.74–4.65
5.39 (2 OH; 4.9), 5.40 (4.8), 5.43 (5.0) 5.43–5.38
HO(3b)
HO(6b)
HO(2c) 5.25 (4.9), 5.28(5.0)
HO(3c) 5.03 (4.6)
HO(4c) 5.00 (5.4), 5.01 (6.0)
HO(6c) 4.62 (5.5), 4.63 (5.5)
4.74 (<1.5), 4.67 (<1.5)
4.72–4.65
5.22 (4.9)
5.00 (4.9)
4.98(5.4)
4.74–4.65
4.74–4.65
5.22 (4.7)
5.01 (4.4)
4.98(5.0)
4.58(5.2)
4.59 (5.3)
a) Broad s, w1/2 =4.7–5.0 Hz.
same chemical shift is not in agreement with an intramolecular H-bond of HO(2a) of T-
x to C(9’)=O, nor is such a H-bond expected, since this C=O group acts already as H-
acceptor of HO(8’) that resonates at the expected low field (12.46–12.50 ppm). A fast
H/H exchange of HO(2a) of T-x is responsible for the broadening of the signals; it is
probably caused by the nearby phenolic HO(8’). The vicinal J(H,OH) of the other
OH groups of the E chain of T-x-x are similar to the corresponding coupling constants
of T-x (DJ mostly smaller than 0.3 Hz). The values of J(2b,OH) and J(2c,OH) (4.7–5.0
Hz), and of J(6b,OH) and J(6c,OH) (5.1–5.2 Hz) of T-x and T-x-x do not hint at per-
sistent intramolecular H-bonds. The larger J(2a,OH) value of T-x-x (x=2, 4, and 8;
5.5–5.9 Hz) and J(6a,OH) values of T-2 and T-2-2 (5.8–6.0 Hz) may point at weakly
persistent intramolecular H-bonds. Since a small J(2,OH) value is expected for intra-
residue O(2)ꢀH···O(3) or O(2)ꢀH···O(1) H-bonded glucopyranosyl species (J<2
Hz [71]), these larger J(2a,OH) values of T-x-x evidence either an electronic influence
of the p-orbitals of the buta-1,3-diynyl substituent (cf. [72]) or weakly persistent inter-
chain H-bonds of HO(2a). That both factors are operating is suggested by J(2,OH) of
the mono- and double-chained glucopyranoside analogues (5.5–5.6 vs. 6.0 Hz [3]).

Helvetica Chimica Acta – Vol. 89 (2006)
691
1
SIMPLE H-NMR experiments [73] with (D )DMSO solutions of T-4, T-4-4, T-8,
6
and T-8-8 did not show any splitting of OH signals upon titrating with D
A
T
E
N
A
H
R
U
G
2
corroborating the absence of strongly persistent intramolecular H-bonds between OH
groups.
The temperature dependence of the OH signals of T-x and T-x-x (x=2, 4, and 8) in
(
D )DMSO was determined from 298to 348K in 10-K intervals. Dd(OH)/DT values
6
are listed in Table 2. Similarly as observed for N-x and N-x-x, a weak temperature
dependence of d(HO(3a)) and d(HO(3b)) values of T-x and T-x-x (x=2, 4; ꢀ2.0 to
ꢀ
2.3 ppb/K) confirms the inter-residue H-bond of HO(3a) and HO(3b) to O(5) of
the adjacent glucopyranosyl moiety. The weak temperature dependence of the
d(HO(8’)) value of T-x (ꢀ1.5 to ꢀ1.9 ppb/K) evidences an intramolecular H-bond of
HO(8’) to C(9’)=O. Dd(OH)/DT Values in the range of ꢀ4.7 to ꢀ7.2 ppb/K for all
other OH groups of T-x and T-x-x evidence more or less completely solvated OH
groups. The DDd(OH)/DT values for corresponding OH groups of T-x and the E
chain of T-x-x are small (ꢂ0.4 ppb/K), except for HOC(2a) with DDd(OH)/DT values
of 0.4–0.7 ppb/K). The slightly higher jDd(HO(2aE)/DTj values of T-x-x may suggest
that HO(2aE) acts as H-acceptor of a weakly persistent inter-chain H-bond.
a
Table 2. Temperature Coefficients Dd(OH)/DT [ppb/K] of T-x and T-x-x in (D )DMSO )
6
T-2
T-4
T-8
T-2-2
T-4-4
T-8-8
E chain, B chain
E chain, B chain
E chain, B chain
HOꢀC(8’)
HO(2a)
HO(3a)
HO(6a)
HO(2b)
HO(3b)
HO(6b)
HO(2c)
HO(3c)
HO(4c)
HO(6c)
ꢀ1.9
ꢀ7.2
ꢀ2.2
ꢀ5.6
–
ꢀ1.9
ꢀ1.9
ꢀ6.0
–
–
–
ꢀ6.8
ꢀ7.9, ꢀ6.6
ꢀ2.0, ꢀ2.3
ꢀ5.6
ꢀ7.2, ꢀ6.2
ꢀ7.4, ꢀ6.4
b
ꢀ2.3
)
ꢀ2.2
ꢀ2.4, ꢀ2.4
b
b
ꢀ5.0
)
ꢀ5.1
)
ꢀ5.8, ꢀ6.1
ꢀ2.3, ꢀ2.2
ꢀ4.7
ꢀ6.0
–
–
–
ꢀ5.8to ꢀ6.0
ꢀ2.2
ꢀ5.8to ꢀ6.2
b
b
–
–
)
)
b
b)
)
ꢀ5.0 to ꢀ5.2
ꢀ5.7
ꢀ6.0
ꢀ6.7
ꢀ6.0
ꢀ5.1
ꢀ5.7
ꢀ5.9
ꢀ5.8
ꢀ5.0
ꢀ5.3
ꢀ6.1, ꢀ6.2
ꢀ6.6, ꢀ6.2
ꢀ5.9, ꢀ6.2
ꢀ5.2, ꢀ5.5
ꢀ5.8
ꢀ6.2
ꢀ6.2
ꢀ5.3
ꢀ6.3
ꢀ6.3
ꢀ5.9
ꢀ5.7
ꢀ5.3
ꢀ5.2
a) Spectra were recorded from 298to 348K in 10-K intervals. b) Not assigned.
H-Bonding of N-x and N-x-x (x=4 and 8) was investigated by analysis of ROESY
7
spectra [4] ). The analysis was restricted to the cross-peaks between HꢀC(1) and OH
signals, as the signals for HꢀC(2) to HꢀC(6) overlapped. Cross-peaks between the
n
n+1
HO(3 ) and HꢀC(1 ) signals (n=1–3 for N-4 and N-4-4, and 1–7 for N-8 and
n
N-8-8) confirme the inter-residue H-bond of HO(3 ). Cross-peaks between the
n
HO(2) and HꢀC(1) signals of the same unit evidence a weakly persistent O(2 )ꢀ
nꢀ1
H···O(6 ) H-bond. Inter-chain H-bonding of N-4-4 and N-8-8 is restricted to unit I;
7)
See there for the differentiation between ꢁtrueꢂ and relayed ROEs, such as TOCSY/ROE and
exchange/ROE.

6
92
Helvetica Chimica Acta – Vol. 89 (2006)
I
a weak inter-chain H-bond between the two HO(2 ) groups is possible, but could not be
unambiguously proven.
ROESY Spectra of T-4, T-4-4, T-8, and T-8-8 were recorded under identical condi-
tions. Signal overlap prevents the assignment of the cross-peaks involving CH groups
other than HꢀC(1). Therefore, the analysis was again limited to interactions between
HꢀC(1) and OH groups, and between OH groups.
The partial ROESY spectrum of T-4 shows seven positive cross-peaks between
4
.0–6.0 ppm (Fig. 4). All HO(2) signals show cross-peaks with the HꢀC(1) signal of
the same residue (peaks #1 to #4). These intra-residue interactions reveal weakly per-
sistent inter-residue H-bonds of HO(2 ) to HOC(6 ), HO(2 ) to HOC(6 ), and
HO(2 ) to HO(6 ) – as also observed for N-4 – and a weakly persistent bifurcated
H-bond of HO(2 ) to the acetylene and C(9’)=O groups (marked in blue in Fig. 5).
IV
III
III
II
II
I
I
III
IV
Additional positive cross-peaks between the signals of HO(3 ) and HꢀC(1 ) (peak
II
III
I
II
#
5), HO(3 ) and HꢀC(1 ) (peak #6), and HO(3 ) and HꢀC(1 ) (peak #7) confirm
the inter-residue H-bonds of HO(3) to O(5) of the next residue (marked in red in
Fig. 5).
Fig. 4. ROESY Spectrum of T-4 : cross-peaks between HꢀC(1) and OH signals
The partial ROESY spectrum of T-4-4 between 4.0–6.0 ppm is depicted in Fig. 6.
I
II
I
I
HꢀC(1 ), HꢀC(1 ), HO(2 ), and HO(3 ) of the B chain resonate at a position that dif-
fers from the one for the corresponding H-atoms of the E chain. The unambiguous
assignment of the signals of T-4-4 is based on the stronger downfield shift of HO(2 )
I
by the buta-1,3-diynyl group [70], and on the analysis of a DQFCOSY and a TOCSY
spectrum. The ROESY spectrum of T-4-4 shows cross-peaks analogous to those
observed for T-4 (peaks #1–#7); they evidence the completely persistent inter-residue

Helvetica Chimica Acta – Vol. 89 (2006)
693
Fig. 5. Interpretation of the ROESY spectrum of T-4 ; arrows indicate ROE cross-peaks and hashed
lines H-bonds
O(3)ꢀH···O(5) H-bonds and weakly persistent inter-residue H-bonds O(2)ꢀH···O(6)
(tautomer A in Fig. 7). The cross-peak pairs #4B/#4E and #7B/#7E reveal only intra-
chain contacts; there are no cross-peaks for the corresponding inter-chain interactions.
IV
An additional cross-peak (#8) is observed between the signals of HꢀC(1 ) and one
I–III
HO(6) signal of the multiplet for HO(6 ). For geometric reasons, it is best assigned
IV
III
to an intra-chain inter-residue close contact between HꢀC(1 ) and HO(6 ). Analo-
III
II
II
gous close contacts between HꢀC(1 ) and HO(6 ), and between HꢀC(1 ) and
I
HO(6 ) may contribute to the cross-peaks #6 and #7. They evidence weakly persistent
inter-residue H-bonds from HO(6) to HO(2) (interactions and H-bonds in green in tau-
tomer B; Fig. 7). It is noteworthy that the cross-peaks between HꢀC(1) and OH signals
reveal inter-residue flip-flop H-bonds between HO(6) and HO(2) of the double-
Fig. 6. ROESY Spectrum of T-4-4: cross-peaks between HꢀC(1) and OH signals

6
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Helvetica Chimica Acta – Vol. 89 (2006)
Fig. 7. Interpretation of the ROESY spectrum of T-4-4; arrows indicate ROE cross-peaks and hashed
lines H-bonds
chained T-4-4 and N-4-4, but not of the mono-chained T-4 and N-4 (only cross-peaks for
unidirected O(2’)ꢀH···O(6)H H-bonds).
There are several positive cross-peaks between OH groups (peaks #9 to #14) in the
ROESY spectrum of T-4-4, whereas the ROESY spectrum of N-4-4 showed only a sin-
III
III
gle positive cross-peak between HO(2 ) and HO(3 ). The inter-residue H-bonds
O(6)ꢀH···O(2’) of tautomer B of T-4-4 suggest a close intra-residue contact of
IV
HO(2) and HO(3). This is confirmed by cross-peaks between the signals of HO(2 )
IV
III
III
IIE
II
and HO(3 ) (#9), HO(2 ) and HO(3 ) (#10), HO(2 ) and HO(3 ) (#11E),
IIB
II
IE
I
IB
HO(2 ) and HO(3 ) (#11B), HO(2 ) and HO(3 ) (#12E), and finally HO(2 ) and
HO(3 ) (#12B); they evidence an intra-residue close contact between HO(2) and
I
HO(3) also for units I (marked with pink arrows in tautomer B). There are two addi-
I
III
tional positive cross-peaks between the broad singlet of HO(3 )/HO(3 ) and, on the
IV
one hand, the triplet of HO(6 ) (#13) and, on the other hand, the doublet of
IV
HO(2 ) (#14). Cross-peak #13 may be assigned to a close intra-chain contact between
IV
IV
III
HO(6 ) (gg conformation for HOC(6 )) and HO(3 ), whereas cross-peak #14 cannot
be assigned to an intra-chain contact (MM3* modeling predicts a H···H distance of 4.75
IV
III
Š
distance between HO(2 ) and HO(3 )). A close inter-chain contact is possible
IVE
IIIB
between HO(2 ) and HO(3 ) but considered improbable since there are no addi-
tional cross-peaks suggesting close contacts between OH groups of the other units,
especially of unit I.

Helvetica Chimica Acta – Vol. 89 (2006)
695
Fig. 8. ROESY Spectrum of T-8 : cross-peaks between HꢀC(1) and OH signals (analogous numbering
of cross-peaks as in T-4-4)
The ROESY spectrum of T-8 (Fig. 8) shows similar cross-peaks as the ROESY
spectrum of T-4-4 for the E chain, confirming the inter-residue H-bond O(3)ꢀ
H···O(5’) and the inter-residue flip-flop H-bond between HO(2) and HO(6’). The
ROESY spectrum of T-8-8 (Fig. 9) shows the same cross-peaks as the ROESY spec-
trum of T-4-4 if one assumes that the cross-peaks #6, #7, #9, and #14 are hidden by
strong negative absorptions. Hence, we expect that T-4-4 and T-8-8 adopt in
(
D )DMSO conformations which avoid inter-chain H-bond interactions. This is feasible
6
in a V-shape arrangement of the cellooligosyl chains, as illustrated by the MM3*-calcu-
lated O(6)ꢀH···O(2’) H-bonded tautomer of T-8-8 in Fig. 10 (considered a model of
cellulose almost completely dissolved in DMSO). This structure suggests at best an
IB
IE
inter-chain H-bond between HO(2 ) and HO(6 ). However, there are no cross-
peaks between these OH signals in the ROESY spectra of T-4-4 and T-8-8.
1
3
4
.2. Comparison of the CP/MAS C-NMR Spectra of T-x and T-x-x (x=1, 2, 4, and
13
8
) with Those of Cellulose I , I , and II. The solid-state CP/MAS C-NMR chemical
shifts of T-x and T-x-x (x=1, 2, 4, and 8) are listed in Table 3 ). The assignment of
the signals is based on a comparison with the data in D O solution (especially of the
a
b
8
AHCTREUNG
2
glucosides T-1 and T-1-1), and with the data of N-x and N-x-x in the solid state. The
8)
We thank Prof. Dr. Beat H. Meier, Ashwin Vorhofen, and Matthias Ernst, Laboratory of Physical
13
Chemistry, ETH Zurich, for the CP/MAS C-NMR spectra of T-x, T-x-x, A-x, and A-x-x.
A
H
R
U
G

6
96
Helvetica Chimica Acta – Vol. 89 (2006)
Fig. 9. ROESY Spectrum of T-8-8: cross-peaks between HꢀC(1) and OH signals (analogous number-
ing of cross-peaks as in T-4-4)
intensity of the anthraquinonyl and ethynyl signals decreases with increasing saccharide
chain length; the spectra of T-8 and T-8-8 essentially show only signals of the central
units b.
13
The solid-state CP/MAS C-NMR spectra of the mono-chained T-x (x=1, 2, 4, and
) are well-resolved (Fig. 11). The spectrum of the glucoside T-1 shows separate peaks
8
for all C-atoms of the glucopyranosyl-ethynyl moiety; the d values are very similar to
those of the solution spectrum in (D )DMSO [3] (Ddꢂ2.0 ppm, with the exception of
6
4
.6 ppm for CꢁCꢀC(1)). Noteworthy are the downfield shift for C(5) resonating at 79.0
ppm, the upfield shift for C(1) appearing at 70.6 ppm, and signals for the ethynyl moiety
at 94.1 and 88.6 ppm. The signals for the ethynyl group and for C(4c) of T-2 and T-4 are
unambiguously assigned by a comparison with the corresponding values of T-1 and by
the decrease of their intensity in T-4. The duplication of the signals for C(1c) and C(4a)
of T2 (107.8/106.4 and 84.6/83.8 ppm, resp.; Table 3) hints at the presence of two differ-
ent molecules in the unit cell. One signal is observed for each C(6a) and C(6c) (62.1/
61.1 ppm). The spectrum of T-4 exhibits the expected three peaks for C(1b) and
C(1c) (109.3, 108.0, and 106.3 ppm), but in a ratio of ca. 3 :2 :1. Similarly, C(4a) and
C(4b) of T-4 give rise to two peaks at 82.4 and 83.0 ppm in a ratio larger than 3 :1,

Helvetica Chimica Acta
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Vol. 89 (2006)
697
Fig. 10. MM3*-Modeled, V-shaped conformer of T-8-8 possessing inter-residue O(6)ꢀH···O(2’) H-
bonds (HꢀC atoms are omitted for enhanced clarity)
whereas C(6a), C(6b), and C(6c) resonate as a single peak at 61.5 ppm with a weak
shoulder at 61.0 ppm. The ratios for the C(1) and C(4) signals again suggest two differ-
ent molecules in the unit cell. There is no downfield shift for C(4a) and C(6) of T-2 and
T-4, and of C(4b) of T-4 ; this clearly evidences that the cellobioside T-2 and the cello-
tetraoside T-4 are not H-bonding models for cellulose II. At best, they mimic cellulose
chains at the surface of the crystallites (see Introduction). In contradistinction, the strik-
13
ing similarity of the solid-state CP/MAS C-NMR spectrum of T-8 and the one of cel-
lulose II, especially the downfield shift of C(4) and C(6) at 89.9/88.7 and 63.8 ppm,
respectively, reveals that the cellooctaoside T-8 is an excellent mimic of cellulose II
in the interior of the crystallite.
13
The solid-state CP/MAS C-NMR spectrum of the C -symmetric diglucoside T-1-1
1
is well-resolved (Fig. 12 and Table 3). The assignment of the peaks of T-1-1 is based on a
comparison with its solution spectrum in (D )DMSO [3] and the solid-state spectrum of
6
13
T-1. The solid-state CP/MAS C-NMR spectra of T-2-2 and T-4-4 show only broad sig-
nals, probably due to a low degree of crystallinity. According to the upfield shift of
C(4b) and C(4c) (<82 ppm), T-2-2 and T-4-4 do not mimic the H-bonds of cellulose
13
I. The solid-state CP/MAS C-NMR spectrum of T-8-8 is much better resolved and

6
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Helvetica Chimica Acta – Vol. 89 (2006)
13
Table 3. CP/MAS C-NMR Chemical Shifts [ppm] of the Mono- and Bis-C-glucoside T-1 and T-1-1, the Mono- and
Bis-C-Cellodextrins T-x and T-x-x (x=2, 4, and 8), and the Cellulose Polymorphs I , I [19], and II [6]
a
b
C(1) of units c
and b
C(4) of units a
and b )
C(2), C(3), C(5), C(1) of
unit a, and C(4) of unit c
C(6)
a
T-1
T-2
–
–
79.0, 76.4, 72.5, 70.6, 68.7
78.5, 77.6, 77.0, 75.6, 74.3,
61.5
62.1, 61.1
107.8, 106.4
84.6, 83.8
7
2.6, 68.8
T-4
109.3, 108.0,
83.0, 82.4
78.5, 77.7, 76.4, 74.4, 72.8,
68.7
61.5
106.3
T-8
Cellulose II
T-1-1
108.3, 106.2
108.3, 106.2
–
89.9, 88.7
89.9, 88.7
–
77.7, 76.0, 73.7
77.8, 75.9, 73.8
79–77, 74.2, 72.9, 71.9,
63.8
64.2, 63.6
59.9, 59.1
6
7.8
T-2-2
T-4-4
T-8-8
Cellulose Ib
Cellulose Ia
104.2 (br.)
103.9 (br.)
107.0, 105.7
107.0, 105.2
106.3
84.0 (br.)
83.5 (br.)
89.9
90.1, 89.3
91.0, 90.2
76.72 (br.), 72.0 (br.)
75.0 (br.)
76.1, 73.0
76.1, 75.3, 73.7, 72.5
75.8, 73.8, 73.0, 72.0
62.7 (br.)
62.8 (br.)
66.3, 64.3
66.8, 66.1
66.5
C(9) and C(10)
of the template
C(8) of the
template of T-x
other signals
of the template
CꢁC and CꢁCꢀCꢁC
T-1
T-2
T-4
T-1-1
T-2-2
T-4-4
185.9, 178.3
187.0, 179.1
186.9, 178.9
184.3, 180.8
180.1 (br.)
163.2
160.1
160.0
–
–
–
140.9, 116.0
144.0, 116.0
143.4, 116.0
143.6, 122.0
141.3, 121.8
143.0, 122.0
94.1, 88.6
92.7, 85.6
92.3, 86.1
94.92, 86.1, 82.0, 80.7, 67.8
92.7 (br.)
93.9 (br.)
a) Except C(4) of T-1 and T-1-1.
resembles that of cellulose I , especially with regard to the two peaks for C(1) at 107.0
b
and 105.7 ppm. A broad peak for C(4) of T-8-8 at 89.9 ppm replaces the double-headed
signal for C(4) of cellulose I at 90.1/89.3 ppm. We conclude that T-8-8, unlike N-8-8, is
b
indeed a H-bond mimic of cellulose I. While a comparison of T-8-8 with N-8-8 strongly
suggests that implementation of the phase shift is crucial to mimic cellulose I, it is not
yet clear which one of the three factors – the rigid linkers, the optimised distance
between the chains, and the phase shift between the chains – is mostly responsible to
render T-8-8 a good mimic of cellulose Ib.
4
.3. Comparison of the X-Ray Powder-Diffraction Spectra of T-8 and T-8-8 with
Those of Celluloses. As several attempts to grow single crystals of T-8 and T-8-8 suitable
for X-ray diffraction failed, we analysed these compounds by X-ray powder diffraction
and compared the spectra with those of celluloses [5] (Fig. 13). The powder-diffraction
9
spectra of T-8-8 and T-8 ) resemble strongly that of cellulose I and cellulose II, respec-
tively, confirming that T-8-8 is an oligomeric mimic of cellulose I and T-8 of cellulose II.
9)
We thank Prof. Dr. Reinhard Nesper and Dr. Qinxing Xie, Laboratory of Inorganic Chemistry, ETH
Zurich, for the powder diffraction spectra of T-8 and T-8-8.

Helvetica Chimica Acta – Vol. 89 (2006)
699
13
Fig. 11. CP/MAS C-NMR Spectra of T-x (x=1, 2, 4, and 8) and cellulose II (from [6])
13
5
. Comparison of the CP/MAS C-NMR Spectra of A-x and A-x-x with Those of
Cellulose Triacetates I and II. Cellulose triacetates I (CTA I) and II (CTA II) are pre-
pared by acetylating native cellulose I and mercerised cellulose II, respectively, and
maintain the polymorphic character of cellulose [74]. Similar to cellulose I and II,
CTA I can be transformed into CTA II, while the reverse process is unknown. Remark-
ably, both CTA I and CTA II were obtained from native celluloses by homogeneous or
heterogeneous acetylation, respectively, while the same conditions transformed cellu-
lose II only into CTA II. Based on the X-ray fiber-diffraction patterns and stereochem-
ical model analysis, a parallel chain orientation was proposed for CTA I [75] and an
antiparallel one for CTA II [76]. The unit cell of CTA I (orthorhombic, P2 ) contains
1
two parallel chains, whereas the unit cell of CTA II (orthorhombic, P2 2 2 ) contains
1
1 1

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Helvetica Chimica Acta – Vol. 89 (2006)
13
Fig. 12. CP/MAS C-NMR Spectra of T-x-x (x=1, 2, 4, and 8), and cellulose I and I (from [6])
a
b
two parallel and two antiparallel chains. However, there are experimental results in
conflict with Spragueꢂs suggested analogy between celluloses and CTAs (see the excel-
lent discussion in [77]) casting some doubt on the unambiguous assignment of the polar-
ity of the chains in CTA I and CTA II.

Helvetica Chimica Acta – Vol. 89 (2006)
701
Fig. 13. X-Ray powder-diffraction patterns of T-8, T-8-8, cellulose I to IV, and amorphous cellulose
13
The CP/MAS C-NMR spectra of CTA I and CTA II show characteristic differen-
13
ces [78]. Analysis of C-enriched samples [79] led to a complete assignment of the sig-
nals for C(2) to C(5). The spectrum of CTA I displays a single set of signals, whereas
CTA II shows a double set of signals for C(2) to C(6) evidencing two different 2,3,6-
tri-O-acetyl-b-D-glucopyranosyl units [77] (Fig. 14). C(1) of CTA I resonates at 102.4
ppm, C(1) of CTA II at 100.4 ppm, C(6) of CTA I at 62.5 ppm, and C(6) of CTA II
at 65.2 and 67.3 ppm (Table 4). The different values for C(6) may indicate a different
orientation of the (acetoxy)methyl group, or the effect of the different environment.
If Horiiꢂs rule [80] should be applicable to acetylated glucopyranosides, then CTA I
addopts the gg, and CTA II the tg and gt conformation.

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Helvetica Chimica Acta – Vol. 89 (2006)
13
Fig. 14. CP/MAS C-NMR Spectra of A-x, A-x-x (x=2, 4, and 8), CTA I, and CTA II (from [78])

Helvetica Chimica Acta – Vol. 89 (2006)
703
13
Table 4. CP/MAS C-NMR Chemical Shifts [ppm] of A-x, A-x-x (x=2, 4, and 8), CTA I [79], and CTA II
[79]
C(1) of units c CꢁC and
C(4) of units
CꢁCꢀCꢁC a and b
C(2), C(3), C(5), C(1) of
unit a, and C(4) of unit c
C(6)
and b
A-2
A-4
103.0
103.1 (2 C),
95.7, 83.6
94.8, 85.0
79.6, 77.6
79.9 (2 C),
77.6
73.3, 72.0, 70.4, 69.6
75.4, 73.8, 72.5, 70.7, 69.1
63.8, 62.9
63.9, 62.1
(3 C)
100.1
A-8
A-2-2
103.4 (br.)
101.3
81.2 (br.)
92.6, 85.6, 78.0 (br.)
4.3, 81.9
76.9 (br.)
73.1, 70.6, 69.9, 68.0
69.9, 66.9
64.8, 61.3
8
A-4-4
A-8-8
CTA I 102.4
CTA II 100.4 (2 C)
103.0 (2 C),
92.2
80.0 (2 C),
78.2
81.6 (br.)
80.3
75.5, 73.8, 72.5, 72.0, 70.7,
69.1, 68.3, 67.4
65.3, 62.4
(3 C)
69.6, 67.0
62.5
99.8
103.4 (br.)
76.7 (br.)
76.2 (2 C), 72.9 )
a
–
–
80.1, 78.7
76.6, 75.8 (2 C), 74.8, 73.5,
67.3, 65.2
a
7
2.5 )
C=O of Ac
Me of Ac C(9) and C(10) of C(8) of the template of A-x Other signals of
the template
the template
A-2
A-4
171.6, 171.1, 22.4, 21.9, 185.4, 181.3
70.5, 169.9 20.9, 19.9
172.0, 170.5, 21.7, 20.4
69.8
162.4
163.3
139.0, 115.0
1
185.7, 180.6
139.0, 115.0
1
A-8
A-2-2
A-4-4
173.5
170.5
24.3
22.0, 20.6
137.7
141.0, 119.0
141.0, 119.0
182.6, 182.0, 179.9 –
172.5, 172.0, 21.8, 20.5
70.8, 169.9
173.6
181.5
–
1
A-8-8
CTA I 171.9, 169.4
24.2
22.0, 20.9
–
–
–
–
CTA II 173.2, 172.2, 23.4, 20.8–
70.0
–
–
1
a) For a complete assignment of these lines, see [77].
13
Surprisingly, according to the studies of the CP/MAS C-NMR and X-ray diffrac-
tion spectra of a series of peracetylated cello-oligomers by VanderHart et al. [78] and
Kono et al. [64][81], the X-ray diffraction spectra of peracetylated cellopentaose and
higher cello-oligosides are similar to those of CTA I. In view of these results, we con-
sidered that analysis of the X-ray diffraction spectra of A-x and A-x-x (x=2, 4, and
8) may contribute to understand the (apparent) opposite chain orientation in the
solid state of free and acetylated higher cellodextrins.
13
The CP/MAS C-NMR spectra of the template-bound peracetylated A-x and A-x-x
x=2, 4, and 8), and of CTA I and CTA II are depicted in Fig. 14, and their chemical
(
shift values are compiled in Table 4. Surprisingly, the spectra of the mono-chained A-
x (especially of the tetraoside and octaoside) resemble the corresponding spectra of
the double-chained A-x-x. C(1) of the tetraosides A-4 and A-4-4 resonates as two
peaks in a 2 :1 ratio at 103.4–103.1 and 101.3–99.8ppm; the former peak is assigned
to C(1b) and the latter one to C(1c). C(6) of A-4 and A-4-4 resonates as two peaks

7
04
Helvetica Chimica Acta – Vol. 89 (2006)
in a 1:3 ratio at 65.3–63.9 and 62.4–62.1 ppm; the weaker peak was assigned to C(6c),
and the stronger one to C(6a) and C(6b). With the exception of the signals for unit c
that find no equivalent in the spectra of CTA I and CTA II, the spectra of A-4 and
A-4-4 resemble the spectrum of CTA I. The (less well resolved) spectra of the cellooc-
taosides A-8 and A-8-8, however, resemble strongly the spectrum of CTA II, particu-
larly when considering the two peaks for C(6), the broad signals for C(4) and for
C(2)/C(3)/C(5), and the single peak for the Me groups. The similarity between the
13
solid-state CP/MAS C-NMR spectra of A-4 and A-4-4 and the one of CTA I, and
of A-8 and A-8-8 with the one of CTA II is opposite to the observations in the cellodex-
trin series. At first view, this result is surprising especially for A-8-8 which possesses par-
allel cellosyl chains. However, the antiparallel orientation of two molecules of A-8-8
leads to two parallel and two antiparallel chains, as required for CTA II. Any further
analysis must resolve the question about the origin of the signal-doubling in the
13
solid-state C-NMR spectrum of CTA II. Is it due to different glucopyranosyl units
in the repeating cellobiosyl moiety of all chains, to different parallel chains, or to differ-
ent antiparallel chains? Better resolved diffraction analyses of CTA II and its oligo-
meric models should allow to answer this fundamental question.
13
6
. Conclusions. Solid-state CP/MAS C-NMR spectroscopy and X-ray powder-dif-
fraction analysis reveal that T-8 and T-8-8 are oligomeric mimics of cellulose II and I ,
b
1
respectively. A combined analysis of the H-NMR data (d(OH), J(H,OH), Dd(OH)/
DT, ROESY spectra) of the mono- and double-chained template-bound cellosides
T-4, T-8, T-4-4, and T-8-8 in (D )DMSO solution reveals strong intra-strand inter-resi-
6
due C(3)OH···O(5’) H-bonds, weakly persistent inter-residue flip-flop H-bonds
between HO(2) and HO(6’), and at best a weakly persistent inter-strand H-bond
I
I
between HO(2 ) of the ethynylated chain E and HO(6 ) of the buta-1,3-diynylated
13
chain B of T-4-4 and T-8-8. The solid-state CP/MAS C-NMR spectra of A-4 and A-
-4 resemble that of CTA I, and those of A-8 and A-8-8 resemble that of CTA II, oppo-
4
site to the findings in the cellodextrin series.
We thank the Swiss National Science Foundation and F. Hoffmann-La Roche AG, Basel, for generous
support, Mrs. Brigitte Brandenberg for recording the 500-MHz NMR spectra, and Prof. Bernhard Jaun for
helpful discussions.
Experimental Part
General. See [3]. Lewis acids such as AgOTf, CdCO , HgCl , Hg(CN) , and BF ·OEt were used
3
2
2
3
2
directly without purification. IR Spectra (ca. 3% soln. in CHCl or CH Cl soln. or 1% KBr pills): Per-
3
2
2
1
13
kin-Elmer 298 and 1600 FT-IR spectrometer. NMR Spectra: Varian XL-300 ( H: 300 MHz, C: 75
MHz), or a Bruker AMX-500 or -600 (500 or 600 MHz, resp.) in deuteriated solvents (CDCl3 and
(
D )DMSO, from Dr. Glaser AG, Basel); chemical shifts d in ppm and couplings constants J in Hz. In
6
1
ambiguous cases, H-assignment is based on selective homonuclear decoupling experiments and 2D
experiments. The CP/MAS C-NMR experiments were performed on a Bruker NMR spectrometer
13
1
13
with 500-MHz resonance frequency for H and 125 MHz for C. The spectrometer is equipped with a
.5-mm MAS double resonance probe from Chemagnetics (Ft. Ciollins, Colorado, USA). The sample
2
spinning frequency was 20 kHz for all samples. The contact time for cross-polarisation is 3 ms and the
1
data-acquisition time is 25 ms. The radio-frequency field strengths are 100 and 84 kHz for the H and
13
the C channel, resp. The numbers of scans collected vary from 10000 to 20000 with a repetition of 8s
for all samples.