Biochemical characterisation and assessment of fibril-forming ability of collagens extracted from Bester sturgeon Huso huso × Acipenser ruthenus

Article abstract of DOI:10.1016/j.foodchem.2014.03.075

Collagens purified from Bester sturgeon organs were characterised biochemically, and their fibril-forming abilities and fibril morphologies formed in vitro clarified. Yields of collagens were 2.1%, 11.9%, 0.4%, 18.1%, 0.4%, 0.8% and 0.03% (collagen dry weight/tissue wet weight) from scales, skin, muscle, swim bladder, digestive tract, notochord and snout cartilage, respectively. Using SDS-PAGE and amino acid composition analyses, collagens from scales, skin, muscle, the swim bladder and digestive tract were characterised as type I, and collagens from the notochord and snout cartilage as type II. Denaturation temperatures of the collagens, measured using circular dichroism, were 29.6, 26.8, 29.0, 32.9, 31.6 and 36.3 °C in scales, skin, muscle, swim bladder, digestive tract, and notochord, respectively. For fibril formation, swim bladder and skin collagen showed a more rapid rate of increase in turbidity, a shorter time to attain the maximum turbidity, and formed thicker fibrils compared with porcine tendon type I collagen.

Full text of DOI:10.1016/j.foodchem.2014.03.075

Food Chemistry 160 (2014) 305–312
Contents lists available at ScienceDirect
Food Chemistry
journal homepage: www.elsevier.com/locate/foodchem
Biochemical characterisation and assessment of fibril-forming ability of
collagens extracted from Bester sturgeon Huso huso  Acipenser ruthenus
⇑
Xi Zhang , Mika Ookawa, Yongkai Tan, Kazuhiro Ura, Shinji Adachi, Yasuaki Takagi
Graduate School of Fisheries Sciences, Hokkaido University, 3-1-1 Minato-cho, Hakodate, Hokkaido 041-8611, Japan
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 24 June 2013
Received in revised form 24 February 2014
Accepted 14 March 2014
Available online 24 March 2014
Collagens purified from Bester sturgeon organs were characterised biochemically, and their fibril-forming
abilities and fibril morphologies formed in vitro clarified. Yields of collagens were 2.1%, 11.9%, 0.4%, 18.1%,
0.4%, 0.8% and 0.03% (collagen dry weight/tissue wet weight) from scales, skin, muscle, swim bladder,
digestive tract, notochord and snout cartilage, respectively. Using SDS–PAGE and amino acid composition
analyses, collagens from scales, skin, muscle, the swim bladder and digestive tract were characterised as
type I, and collagens from the notochord and snout cartilage as type II. Denaturation temperatures of the
collagens, measured using circular dichroism, were 29.6, 26.8, 29.0, 32.9, 31.6 and 36.3 °C in scales, skin,
muscle, swim bladder, digestive tract, and notochord, respectively. For fibril formation, swim bladder and
skin collagen showed a more rapid rate of increase in turbidity, a shorter time to attain the maximum
turbidity, and formed thicker fibrils compared with porcine tendon type I collagen.
Keywords:
Type I collagen
Type II collagen
Denaturation temperature
Notochord
Ó 2014 Elsevier Ltd. All rights reserved.
SEM
1. Introduction
2009; Kittiphattanabawon, Benjakul, Visessanguan, Nagai, &
Tanaka, 2005; Nagai & Suzuki, 2000; Singh, Benjakul, Maqsood, &
Kishimura, 2011; Zhang et al., 2009).
Collagen, in the form of elongated fibrils, contributes to the
physiological functions and mechanical properties of skin, tendon,
bone, cartilage and other tissues (Ikoma, Kobayashi, Tanaka, Walsh,
& Mann, 2003). Collagen has been widely used in many industries
such as the food, photographic film, cosmetic, and leather indus-
tries (Zhang, Liu, & Li, 2009). Furthermore, because of its low anti-
genic activity, high cell adhesion properties, biocompatibility, and
biodegradability, collagen-based biomaterials have been applied in
medical research, as well as tissue engineering (Parenteau-Bareil,
Gauvin, & Berthod, 2010). The most common sources of collagen
for biomaterials and tissue engineering are bovine skin and
tendons, porcine skin, and rat tail (Parenteau-Bareil et al., 2010).
However, concerns about the use of land animal collagens are
growing because of the risk of common human diseases. Religious
beliefs also restrict usage of porcine or bovine collagens in certain
regions in the world. In recent years, as an alternative source to
porcine or bovine collagen, fish collagen has received increasing
attention, for its advantages of using highly abundant fish offal,
and avoiding human diseases and religious objections (Jongjareon-
rak, Benjakul, Visessanguan, & Tanaka, 2005a). The biochemical
nature of collagens extracted from fish skin, scales, and bones have
been reported in many studies (Duan, Zhang, Du, Yao, & Konno,
Sturgeon is highly valued as a food fish, and is especially famous
for its caviar. Contemporary sturgeon culture was begun in the
1980s, and worldwide production had increased to ca. 25,000 t
by 2008 (Zhang, Shimoda, Ura, Adachi, & Takagi, 2012). Thus, stur-
geon culture now forms the basis of an important and promising
industry. However, the culture cost of sturgeon is higher than other
fishes, because caviar production requires long culture times. On
the other hand, the lack of utilisation of other parts of sturgeon
constrains the development of its culture. Therefore, we believe
that if skin, offal and cartilage of sturgeon are used as a source of
collagen, the value of the by-products would increase, and waste
disposal problems could be alleviated.
Zeng et al. (2012) have reported the structure and characteris-
tics of collagen extracted from cobia Rachycentron canadum skin
for the utilisation in biomaterials, which should increase the value
of cobia offal. However, the important collagen property, the fibril-
forming ability, and morphology for biomaterial utilisation has not
been reported. The collagen molecule is distinct from other pro-
teins as its three polypeptide chains (a-chains) form a unique
triple helical structure. The molecules self-assemble into fibrils
in vitro when collagen solution is adjusted to an appropriate
temperature, pH, and ionic strength (Kadler, Holmes, Trotter, &
Chapman, 1996). After fibril formation, the low denaturation tem-
perature of fish collagen can be improved to more than 40 °C, and
⇑
Corresponding author. Tel./fax: +81 138 40 5551.
E-mail address: ocean_zhangxi@yahoo.co.jp (X. Zhang).
http://dx.doi.org/10.1016/j.foodchem.2014.03.075
0308-8146/Ó 2014 Elsevier Ltd. All rights reserved.

306
X. Zhang et al. / Food Chemistry 160 (2014) 305–312
therefore it is possible for use as a biomaterial for humans (Bae
et al., 2009). However, there are few studies about the fibril-form-
ing ability of fish collagens.
to estimate the molecular weight. Quantitative analysis of band
intensity was determined by AE-6932GXES Printgraph (ATTO Co.,
Tokyo, Japan) with CS Analyzer ver 3.0 (image analysis system).
Bester, a hybrid sturgeon of Huso huso  Acipenser ruthenus, is
suited to aquaculture because of its rapid growth, ability to grow
in freshwater ponds, and the relative ease of inducing sexual mat-
uration (Zhang et al., 2012). This fish is now widely cultured in Chi-
na (Wei, Zou, Li, & Li, 2011) and Japan. The objective of this study
was to extract and assess the biochemical nature of the collagens
from the Bester sturgeon. In addition, the fibril-forming ability
and morphology of fibrils formed in vitro were investigated.
2.3. Amino acid analysis
Samples were hydrolysed in 6 N HCl at 110 °C for 24 h. The
hydrolysates were evaporated, and the remaining materials
dissolved in citric acid buffer solution, and then analysed using
an automated amino acid analyzer (JLC-500 V, JEOL Ltd., Tokyo,
Japan). Samples were assayed three times and the averages were
used to obtain amino acid compositions. In this study, the number
of cysteine residues was obtained by first determining the number
of cystine residues (a dimeric amino acid) then calculating cysteine
from cystine. The number of tryptophan residues was not
determined.
2. Materials and methods
2.1. Extraction of collagens
A live cultured Bester sturgeon (length = 0.76 m, 2.00 kg) was
procured from the Nanae Fresh-Water Laboratory, Field Science
centre for Northern Biosphere, Hokkaido University, Japan. The fish
was deeply anaesthetized in 2-phenoxyethanol solution and gut-
ted. Scales, skin, muscle, swim bladder, digestive tract, notochord
and snout cartilage were removed and washed with chilled tap
water, then stored at À30 °C until use. Samples were washed with
cold distilled water (4 °C) and cut into small pieces (0.5 Â 0.5 cm).
Skin fat was removed over 24 h in 99.5% ethanol (three solution
changes) with a sample/solution ratio of 1:10. Scales and snout
cartilage samples were removed, non-collagenous proteins ex-
tracted in 0.1 M NaOH with a sample/solution ratio of 1:10 for
6 h, and then samples were decalcified in 0.5 M EDTA with a sam-
ple/solution ratio of 1:10 for 24 h. After that, skin, scales and carti-
lage samples were washed with cold water for 24 h. To extract
collagens, the samples were stirred continuously in a solution of
HCl (pH 2.0) containing 0.1% (w/v) porcine pepsin (EC 3.4.23.1,
1:10,000, Wako Pure Chemical Industries Ltd., Osaka, Japan) with
a sample/solvent ratio of 1:10 (w/v) for 48 h at 10 °C. The mixtures
were centrifuged at 2000g for 1 h to get supernatants, and the res-
idue was re-extracted in the same conditions. The supernatants
were sequentially filtered with membrane filters with pore sizes
2.4. Circular dichroism (CD) measurement
CD spectra were measured using a JASCO model 725 spectrom-
eter (JASCO, Tokyo, Japan). The measurement was performed
according to the method of Ikoma et al. (2003). Lyophilized colla-
gens were dissolved in an HCl solution (pH 3.0) to 1 mg/ml, and
placed into a quartz cell. CD spectra were measured at 190–
250 nm wavelengths at 10 °C under a scan speed of 50 nm/min
with an interval of 0.1 nm. Then a rotatory angle at a fixed wave-
length of 221 nm was measured at 10–50 °C with a rate of 1 °C/
min to determine the collagen denaturation temperature (Td).
2.5. Collagen fibril formation in vitro
Collagen molecules self-assemble into fibrils in vitro (fibril for-
mation) when collagen solution is adjusted to an appropriate tem-
perature, pH, and ionic strength. Fibril formation of collagen from
the skin and swim bladder was performed according to the method
of Bae et al. (2009). Lyophilized collagens were dissolved in an HCl
solution (pH 3.0) to 0.3% (w/v). The collagen solution was mixed
with 0.1 M Na-phosphate buffer (pH 7.4) containing NaCl at 0, 70
and 140 mM. The ratio of the collagen solution/Na-phosphate buf-
fer was 1:2 (v/v), and the final pH of the solution was 7.4. Porcine
tendon collagen (Cellmatrix Type I-A, 0.3%, Nitta Gelatin Inc., Osa-
ka, Japan) was used as a control. With fibril formation, the trans-
parent solution would become white. The mixed solution was
placed into a cell, and the resulting fibril formation at 21 1 °C
was monitored by the absorbance at 320 nm as the turbidity
change using a spectral monitor GeneQuant pro (GE Healthcare Life
Sciences, Tokyo, Japan). This is the measurement on the speed of
collagen fibril formation in a short time.
of 3.0, 0.8, and 0.47 lm (Advantec, Tokyo, Japan). Collagen in the
filtrate was precipitated by adding NaCl to a final concentration
of 1 M. The resulting precipitate was collected by centrifugation
at 2000g at 4 °C for 90 min, and dissolved in an HCl solution (pH
2.0). This process was repeated three times to purify the collagen.
The purified collagen was dialyzed against 50 volumes of distilled
water at 4 °C for 24 h with two changes of water. The dialysate was
lyophilized by freeze dryer (FDU-830, Tokyo Rikakikai Co., Ltd., To-
kyo, Japan). The percentage of dry weight of collagen extracted in
comparison with the wet weight of the initial tissues was calcu-
lated as the collagen yield.
2.2. Sodium dodecyl sulphate polyacrylamide gel electrophoresis
(SDS–PAGE) analysis
2.6. Measurement of degree of collagen fibril formation
SDS–PAGE was performed according to the method of Laemmli
(1970). The lyophilized collagens were dissolved in HCl (pH 3.0,
3 mg collagen/ml), and then mixed at a ratio of 1:2 (v/v) with
the sample buffer (0.5 M Tris–HCl buffer, pH 6.8, with 4% SDS
and 20% glycerol) containing 10% b-mercaptoethanol. The mixed
solution was boiled for 5 min. Ten micrograms of protein were
loaded in each lane. Electrophoresis was performed at 15 mA for
the stacking gel and 20 mA for the 7.5% running gel. After electro-
phoresis, the gel was stained for 30 min using 0.1% Coomassie Bril-
liant Blue R250 solution and destained with a mixture of 20%
ethanol, 5% acetic acid and 2.5% glycerin. Precision plus protein
standards (Bio-Rad Laboratories, Inc., Hercules, CA, USA) were used
Skin and swim bladder collagen fibrils as well as porcine tendon
collagen fibrils were formed for 24 h at 21 1 °C, which was
assumed to be the endpoint of fibril formation, using the same con-
ditions as described above. Then the sample solution was centri-
fuged at 20,000g for 20 min to precipitate fibrils, and the protein
content of the supernatant was measured based on the method
of Lowry (Lowry, Rosebrough, Farr, & Randall, 1951) using bovine
serum albumin as a standard. Degree of collagen fibril formation
was defined as the percentage of the decrease of the collagen con-
centration in the solution after the experiment, which means the
percent of collagen molecules that formed the fibrils.

X. Zhang et al. / Food Chemistry 160 (2014) 305–312
307
2.7. Electron microscope observations
(Jongjareonrak et al., 2005b). Thus pepsin-digestion extracts
collagen molecules without cross-links at the triple helical region.
Therefore, the more rapid extraction of collagen from the swim
bladder under pepsin digestion suggests that inter-molecular
cross-links at the triple helical region of the collagen molecules
in this tissue are likely to be fewer than those in other organs.
The microstructure of skin and swim bladder collagen fibrils
was observed using
a scanning electron microscope (SEM;
JSM6010LA, JEOL Ltd., Tokyo, Japan). The collagen fibrils were
formed using the same conditions as described above for 1 and
24 h at 21 1 °C. The sample suspension of skin and swim bladder
collagens was centrifuged at 20,000g for 20 min to get precipitates
of collagen fibrils. The collagen fibrils were fixed with 2.5% (v/v)
glutaraldehyde in 0.1 M phosphate buffer (pH 7.6) for 3 h at room
temperature, and then rinsed with the phosphate buffer. The fibrils
were sequentially soaked in 70%, 80%, 90%, 95% and 100% ethanol
solutions for 30 min to dehydration, and then in two 30-min
changes of t-butyl alcohol solution. Finally, collagen fibrils were
freeze-dried in t-butyl alcohol solution with a freeze-drying device
(JFD-320; JEOL Ltd.) and coated with gold-platinum using an auto
fine coater (JFC-1600; JEOL Ltd.). For comparison, porcine tendon
type I collagen (Cellmatrix Type I-A, Nitta Gelatin Inc., Osaka,
Japan) was treated similarly and observed.
3.2. SDS–PAGE
The SDS–PAGE patterns of collagens are shown in Fig. 1.
Collagen from scales, skin, muscle, swim bladder and digestive
tract consisted of two
constituents, and the density of the 120-kDa band (
higher than that of the 100-kDa band ( 2-chain), at a ratio of
a-chains (ca. 120 and 100 kDa) as the major
a1-chain) was
a
approximately 2:1 (scale 2.1:1, skin 2.1:1, muscle 2.0:1, swim
bladder 2.0:1, digestive tract 2.1:1). The molecular weight of these
bands, as well as the band pattern, suggest that these collagens are
most likely to be classified as type I collagen. On the other hand,
the ratio which is little different from 2:1 might show the possibil-
ity of the presence of other type collagen in the extracted collagens,
such as the type III collagen which is present in skin and intestines
and the type V collagen which is present in tissues where type I
collagen is expressed (Parenteau-Bareil et al., 2010; Wang et al.,
3. Results and discussion
3.1. Yields of collagens
2014). In addition, the a3-chain has been reported to exist in skin
type I collagen of white sturgeon, Bester sturgeon, and many other
teleost fish (Kimura, 1992). However, it could not be determined
Yields of collagens are summarised in Table 1. The yield of swim
bladder collagen (18.1% on a wet weight basis, 37.7% on a dry
weight basis) was the highest on a percent basis, and snout carti-
lage (0.03% and 0.2%, on a wet and dry weight basis, respectively)
was the lowest. The amount of collagen obtained from one fish
(length = 0.76 m, 2.00 kg) was highest from the skin (6010 mg)
and the swim bladder (4400 mg). The yield of collagen from stur-
geon skin (11.9%, wet weight basis) was therefore higher than that
from bigeye snapper skin (7.5%; Jongjareonrak et al., 2005a), and
similar to skin yields from brownstripe red snapper (13.7%; Jong-
jareonrak, Benjakul, Visessanguan, Nagai, & Tanaka, 2005b) and
striped catfish (12.8%; Singh et al., 2011). The yield of collagen
from sturgeon skin (34.1%, dry weight basis) was higher than that
from black drum skin (18.1%), but lower than that from largefin
longbarbel catfish skin (44.8%) and ocellate puffer fish skin
(55.4%) (Nagai, Araki, & Suzuki, 2002; Zhang et al., 2009). The com-
parable high yield of skin and swim bladder collagen suggests the
possibility for industrial production.
whether the
cause the migration similarity of the
separation of the former from the latter using SDS–PAGE (Kimura,
1992). To determine whether these collagens contain the 3-chain,
a3-chain existed in the collagens in this study, be-
a
3 and 1 chains prevents
a
a
CM-cellulose chromatography should be used (Kimura & Ohno,
1987). Notochord collagen and snout cartilage collagen of the
Bester sturgeon consisted of only one
uent (Fig. 1), which may be classified as type II collagen, as type II
collagen comprises three identical chains (Foegeding, Lanier, &
Hultin, 1996; Kittiphattanabawon, Benjakul, Visessanguan,
a-chain as the major constit-
a
&
Shahidi, 2010). Miller and Mathews (1974) also reported the
collagen from notochord and cartilage of sturgeon is comprised
of a single type of
higher vertebrates.
a-chain, which was similar to a1 (II) chain of
In the collagen extraction process, the solubility of swim blad-
der collagen was much higher than that of others. The collagen
molecule has a unique triple helical structure, and the two termi-
nal ends are non-helical parts known as the telopeptide region.
Covalent bonds through the condensation of aldehyde groups at
the telopeptide region, as well as the triple helical region, form
an inter-molecular cross-linked structure, leading to a decrease
in the solubility of collagen (Duan et al., 2009; Singh et al., 2011).
Pepsin has been reported to cleave peptides in the telopeptide
region without damaging the integrity of the triple helix structure
3.3. Amino acid composition
Amino acid compositions of Bester collagens are shown in
Table 2. All collagens had glycine as the major amino acid, and
were also rich in alanine, proline and hydroxyproline. Generally,
glycine in collagen represents approximately one third of the total
residues and occurs as every third residue in collagen molecules,
except for the telopeptide regions (first 14 amino acid residues
from the N-terminus and the last 10 of the C-terminus) (Singh
et al., 2011; Zhang et al., 2009).
For type II collagen of the notochord and snout cartilage, the
glutamic acid, leucine, hydroxylysine, and imino acid (pro-
line+hydroxyproline) contents were higher than those of type I col-
lagens, and yet the contents of serine, alanine, and lysine were
lower (Table 2). In addition, the degree of hydroxylation of lysine
was calculated to be 62% in notochord collagen and 56% in snout
cartilage (Table 3), which was much higher than those in type I
collagens such as scale (25%) and skin (29%) collagens. Seyer and
Vinson (1974) reported that the high degree of lysine hydroxyl-
ation was one of the criteria to identify type II collagen from type
I collagen. Therefore, notochord collagen and snout cartilage colla-
gen are thought to be type II collagen, in accordance with the result
of SDS–PAGE.
Table 1
Yields (%, based on the wet and dry weight of initial samples) and dry weight (mg) of
collagens purified from Bester sturgeon (0.76 m, 2.00 kg).
Samples
Yields/wet (%)
Yields/dry (%)
Yields (mg)
Scale
Skin
Muscle
Swim bladder
Digestive tract
Notochord
Snout cartilage
2.1
11.9
0.4
18.1
0.4
0.8
3.0
34.1
2.0
37.7
2.8
5.1
80
6010
692
4400
105
460
2
0.03
0.2

308
X. Zhang et al. / Food Chemistry 160 (2014) 305–312
M
1
2
3
4
5
6
7
250 kDa
150 kDa
α1(I)
α1(II)
100 kDa
75 kDa
α2(I)
Fig. 1. SDS–PAGE of collagens from Bester sturgeon. M, high molecular weight marker; lane 1: scale collagen; lane 2: skin collagen; lane 3: muscle collagen; lane 4: swim
bladder collagen; lane 5: digestive tract collagen; lane 6: notochord collagen; lane 7: snout cartilage collagen.
Table 2
Amino acid composition of collagens from Bester sturgeon (expressed as residues/1000 total amino acid residues).
Scale
Skin
Muscle
Swim bladder
Digestive tract
Notochord
Snout cartilage
Asp
Thr
Ser
Glx
Gly
Ala
Cys/2
Val
Met
Ile
48
23
42
76
340
116
2
17
9
11
17
2
47
24
43
69
344
121
2
17
9
11
17
1
48
25
42
78
330
118
2
19
9
13
19
2
45
26
42
77
335
121
1
17
10
12
16
2
48
23
43
76
340
110
1
17
10
11
19
2
48
21
31
96
322
95
1
16
8
11
29
2
48
20
34
86
333
102
1
16
8
10
26
2
Leu
Tyr
Phe
HyLys
Lys
11
9
27
4
12
10
25
4
13
10
26
4
12
13
23
4
12
13
22
4
13
24
15
5
12
22
17
5
His
Arg
Hypro
Pro
53
76
117
193
52
78
114
192
51
80
111
191
54
88
102
190
52
82
115
197
51
81
131
212
52
87
119
206
Imino acid
Table 3
Hydroxylation degree of lysine residues of collagens from the Bester sturgeon (expressed as residues/1000 total amino acid residues).
Scale
Skin
Muscle
Swim bladder
Digestive tract
Notochord
Snout cartilage
Lys
HyLys
27
9
25
10
26
10
23
13
22
13
15
24
17
22
Hydroxylation
25%
29%
28%
36%
37%
62%
56%
3.4. Thermal stability
26.8 °C, muscle collagen 29.0 °C, swim bladder collagen 32.9 °C,
digestive tract collagen 31.6 °C, and notochord collagen 36.3 °C.
This suggests that the inside tissue possesses a higher Td. In the
same species, type II collagen seems to have the higher Td than
type I collagen (Amudeswari, Liang, & Chakrabarti, 1987). Thus,
the highest Td in the notochord collagen may be attributed to its
collagen type (type II). However, for type I collagen, the reason
why inside tissue has a higher Td is not known. We could not mea-
sure Td of snout cartilage because the amount of purified collagen
from this organ was too small. The Tds of type I collagens in fish
species are positively correlated with their preferred environmen-
tal water temperature (Singh et al., 2011).
CD spectra of the collagens from the scales, skin, muscle, swim
bladder, digestive tract and notochord of Bester sturgeon are
shown in Fig. 2a. All collagens showed a rotatory maximum at
221 nm and a crossover point (zero rotation) at about 212 nm,
which are typical characteristics of the collagen triple helical con-
formation (Ikoma et al., 2003). Mensuration of collagen denatur-
ation temperature is shown in Fig. 2b. When the temperature
rose, the CD [221] values decreased due to the decomposition of
the collagen triple helical structure. Denaturation temperature
(Td) was determined as the temperature with the fastest decreased
speed of the CD [221] value. Denaturation temperatures of Bester
sturgeon were as follows: scale collagen 29.6 °C, skin collagen
The Td of notochord collagen was higher than some tropical fish
type I collagens, as described above, and similar to those of

X. Zhang et al. / Food Chemistry 160 (2014) 305–312
309
NaCl 0 mM
(a)
2.2
2.0
1.8
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
150
100
50
221nm
skin
swim bladder
porcine collagen
0
190
200
210
220
230
240
250
-50
scale
-100
-150
-200
-250
-300
-350
-400
skin
muscle
swim bladder
digestive tract
notochord
0
10
20
30
40
50
Time (min)
NaCl 70 mM
2.2
2.0
1.8
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
0
wavelength (nm)
(b)
100
80
60
40
20
0
scale
skin
muscle
swim bladder
digestive tract
notochord
10
20
30
40
50
Time (min)
NaCl 140 mM
2.2
2.0
1.8
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
10
20
30
40
50
-20
Temperature (℃)
Fig. 2. CD spectra (a) and temperature effect on the CD spectra at 221 nm, (b) of the
collagens from scale, skin, muscle, swim bladder, digestive tract and notochord of
Bester sturgeon.
0
10
20
30
40
50
collagens in brownbanded bamboo shark and blacktip shark carti-
lages (34.56–36.73 °C), which were suggested to be a mixture of
type I and II collagens (Kittiphattanabawon et al., 2010). However,
the Td of notochord collagen was lower than calf skin type I colla-
gen (40.8 °C) (Duan et al., 2009) and chick sternal cartilage type II
collagen (43.8 °C) (Cao & Xu, 2008). Differences in Tds amongst
collagens of different species were reported to be correlated to
the different imino acid contents (proline and hydroxyproline)
(Kittiphattanabawon et al., 2005). The mechanism is assumed as
proline and hydroxyproline rich zones of collagen molecules that
are most likely involved in the formation of junction zones that
help to strengthen the triple helix structure (Singh et al., 2011).
Time (min)
Fig. 3. Effects of NaCl concentrations on the in vitro progress of fibril formation of
skin and swim bladder collagen of Bester sturgeon and porcine collagen.
of fibril formation in swim bladder collagen, which suggests a high
speed of nucleation when compared with skin collagen. Addition-
ally, swim bladder and skin collagen both showed a rapid rise in
turbidity compared with porcine tendon type I collagen. Thus, it
can be concluded that sturgeon type I collagen had a higher ability
of fibrillogenesis than porcine collagen in the present experimental
conditions.
3.5. Process of collagen fibril formation in vitro
Compared with the fibril formation rate of porcine tendon col-
lagen, those of swim bladder and skin collagen increased with
increasing ionic strength by the addition of NaCl (Fig. 3), which
suggests that NaCl concentration had a more pronounced effect
on the process of collagen fibrillogenesis in sturgeon. In contrast,
the rates of fibril formation of red stingray skin type I collagen
and salmon skin type I collagen were suppressed by high NaCl con-
centrations (Bae et al., 2009; Yunoki, Nagai, Suzuki, & Munekata,
2004). Different characteristics of collagen fibril formation may
be explained in part by species differences. In addition, they may
also be affected by the method of purification of the collagen, the
solvent system employed in the fibrillogenesis experiment, and
the condition under which fibril formation is initiated (Williams,
Gelman, Poppke, & Piez, 1978). Fibrillogenesis appears to be an
Collagen molecules have the ability to self-assemble from solu-
tion, and it is now generally accepted that all of the information
needed for fibril formation is contained in the collagen molecules
themselves (Helseth & Veis, 1981). In this study, the fibril-forming
ability of skin and swim bladder type I collagen was studied, as the
yields of these two collagens were much higher than others. There-
fore, the production of collagen from these tissues on an industrial
scale may be possible. Fibril formation was monitored by the in-
crease in turbidity at 320 nm (Fig. 3). For type I collagen, swim
bladder collagen showed a more rapid rate of increase in turbidity,
and a shorter time to attain the maximum turbidity than skin
collagen at the same NaCl concentration. There was no lag time

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X. Zhang et al. / Food Chemistry 160 (2014) 305–312
100
95
90
85
80
75
70
3.6. Degree of collagen fibril formation
The degrees of collagen fibril formation of Bester sturgeon were
assessed after 24 h of fibrillogenesis, and the results are shown in
Fig. 4. The degree of skin collagen fibril formation ranged from
91% to 93%, and was slightly increased by increasing NaCl concen-
tration. The degree of swim bladder collagen fibril formation ran-
ged from 97% to 98%, was higher than that of skin collagen and
porcine collagen, and also slightly increased by increasing NaCl
concentration. In a study of red stingray collagen, the degree of fi-
bril formation was also slightly increased with increasing NaCl
concentration (Bae et al., 2009). In contrast to these collagens,
the degree of salmon skin collagen fibril formation was decreased
by increasing NaCl concentration (Yunoki et al., 2004). Taken to-
gether, the results from sturgeon collagen and the reported results
from stingray and salmon skin reveal that the degree of collagen fi-
bril formation varies greatly depending on the source of collagen.
NaCl 0 mM
NaCl 70 mM
NaCl 140 mM
skin
swim bladder porcine collagen
Fig. 4. Effects of NaCl concentrations on degree of fibril formation of skin collagen
and swim bladder collagen of Bester sturgeon and porcine collagen. Error bars show
standard deviation.
entropy-driven self-assembly process, which is driven by the loss
of solvent molecules from the surface of protein molecules and re-
sult in assemblies with a circular cross-section, which minimizes
the surface area/volume ratio of the final assembly (Kadler et al.,
1996). As different NaCl concentration resulted in the different io-
nic strength of solution which may affect the assembly process of
collagen molecules, leading to different speed of collagen forma-
tion. On the other hand, in the lag phase, during which no measur-
able change occurs in turbidimetric analysis, was interpreted as
collagen self-assembly involved in a nucleation mechanism analo-
gous to the formation of crystals from a supersaturated inorganic
salt solution (Helseth & Veis, 1981). The general principles of colla-
gen fibril formation are accepted, yet molecular mechanism of the
assembly process is less known.
3.7. SEM observation of collagen fibrils
SEM images of skin, swim bladder, and porcine tendon collagen
fibrils after 1 h of incubation are illustrated in Fig. 5. The unordered
structural appearance of fibrils in sturgeon collagen indicates that
fibrillogenesis occurs in all collagens and solutions. Sturgeon colla-
gen showed a similar fibril character, and the fibrils seemed to
become thicker with increasing NaCl concentration. For porcine
collagen, only a few fibrils were observed in the images at
140 mM NaCl (e.g. Fig. 5i), suggesting a much lower fibril-forming
ability of porcine collagen than sturgeon collagen in a short time.
SEM images of skin collagen, swim bladder collagen and porcine
tendon collagen fibrils after 24 h incubation are illustrated in Fig. 6.
Fibrils of skin collagen showed an unordered structure and were
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
(i )
Fig. 5. Scanning electron micrographs of fibrils formed at 21 1 °C for 1 h. (a–c) sturgeon skin with NaCl concentration of 0, 70 and 140 mM; (d–f) sturgeon swim bladder
with NaCl concentration of 0, 70 and 140 mM; (g–i) porcine collagen with NaCl concentration of 0, 70 and 140 mM. Scale bars, 5 m.
l

X. Zhang et al. / Food Chemistry 160 (2014) 305–312
311
(a)
(d)
(g)
(b)
(c)
(f)
(i)
(e)
(h)
Fig. 6. Scanning electron micrographs of fibrils formed at 21 1 °C for 24 h. (a–c) sturgeon skin with NaCl concentration of 0, 70 and 140 mM; (d–f) sturgeon swim bladder
with NaCl concentration of 0, 70 and 140 mM; (g–i) porcine collagen with NaCl concentration of 0, 70 and 140 mM. Scale bars, 5 m.
l
thicker than the fibrils from 1 h incubation (Fig. 6a–c). For swim
bladder collagen, some fusiform structures characterised by a
Njieha, & Prockop, 1982). However, there are no data on the effects
of NaCl concentrations on the morphology of fibrils. Similarly, for
fish fibrils, there are a few morphological studies only, such as on
type I collagen from salmon skin (Yunoki et al., 2004), shark skin
(Nomura, Toki, Ishii, & Shirai, 2000) and red stingray skin (Bae
et al., 2009), and no data on the effects of NaCl concentrations on
the morphology of formed fibrils. Therefore, this study is possibly
the first to clarify the effects of NaCl concentrations on the detailed
morphology of fibrils of fish type I collagen formed in vitro. More-
over, to the best of our knowledge, this is the first report about the
fusiform structure of collagen fibrils. The effect of NaCl and/or ionic
strength on the morphology of collagen fibrils formed in vitro is
worthy of study when it is intended to apply collagens for medical
purposes such as a cell culture system and tissue engineering,
because collagen morphology of the scaffold will affect cellular
responses. Further assessments on the biomechanical strength of
collagen gels, as well as cellular responses against the gels, are
needed to industrialise biomaterials made of sturgeon collagens.
transverse periodic banding pattern (one unit width ꢀ0.45
lm)
were observed (Fig. 6e). When NaCl concentration was increased,
more fibrils formed the bigger fusiform structures. At 140 mM
NaCl, almost all the fibrils formed bigger fusiform structures, and
the average width of a unit’s banding pattern became narrower
(one unit width ꢀ0.22
lm) (Fig. 6f). In porcine collagen, a few
fibrils only were observed at 0 mM NaCl (Fig. 6g). At 70 and
140 mM NaCl (Fig. 6h and i), the fibrils were obvious. However,
the fibrils were more slender than those from skin and swim
bladder collagens. Thus, sturgeon type I collagens may have a high-
er fibril-forming ability than type I porcine collagen.
In sturgeon skin and swim bladder collagen, the effect of
increasing NaCl concentration on fibril formation is an increase
in the thickness of the fibrils. In swim bladder collagen, increasing
NaCl concentration promotes a characteristic fusiform structure
after 24 h incubation. The formation process of the fusiform struc-
ture was not clarified in the present experiments. However, some
SEM images appeared to show the fusion of several fibrils into a
fusiform fibril. Therefore, our working hypothesis is as follows:
first, collagen is assembled to form fibrils, then several fibrils are
fused into one fusiform structure. The reason for the construction
of the banding pattern is unknown. The fact that a fusiform struc-
ture was observed only after 24 h incubation suggests that fusion
of collagen fibrils takes longer than fibril formation itself. Although
the reason why only swim bladder collagen has the ability to form
a fusiform structure is not clear, it may relate to the ability of rapid
fibril formation in this collagen.
4. Conclusion
This study assessed the biochemical nature of collagens purified
from the Bester sturgeon H. huso  A. ruthenus. In addition, details
of the fibril-forming ability and morphology of fibrils formed
in vitro were clarified. A quantity of collagens could be extracted
from skin, swim bladder and the notochord of Bester sturgeon. Col-
lagens from scales, skin, muscle, the swim bladder and digestive
tract were characterised as type I collagen, and collagens from
the notochord and snout cartilage characterised as type II collagen.
Sturgeon collagens have two characteristic features: relatively
higher thermal stabilities compared with other fish species, and a
better fibril-forming ability in skin and swim bladder type I
There are some studies in terrestrial vertebrates on the mor-
phology of collagen fibrils formed in vitro, such as type I collagens
from calf skin (Holmes, Capaldi, & Chapman, 1986), rat tail tendon
(Williams et al., 1978), and chick embryo tendon (Miyahara,

312
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Kittiphattanabawon, P., Benjakul, S., Visessanguan, W., Nagai, T., & Tanaka, M.
(2005). Characterisation of acid-soluble collagen from skin and bone of bigeye
snapper (Priacanthus tayenus). Food Chemistry, 89, 363–372.
collagen compared with mammalian type I collagen. The big fusi-
form fibril structures of swim bladder collagen suggest its special
utility for enhancing mechanical strength of collagen-based bioma-
terials. Such high thermal stability and high ability of fibril forma-
tion will open a new field of industrial utilisation for sturgeon
collagens as biomaterials, which may increase the economic value
of sturgeon and accelerate aquaculture development of this fish.
Kittiphattanabawon, P., Benjakul, S., Visessanguan, W.,
& Shahidi, F. (2010).
Isolation and characterization of collagen from the cartilages of brownbanded
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Lowry, O. H., Rosebrough, N. J., Farr, A. L.,
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Acknowledgement
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This study was partly supported by the Regional Innovation
Strategy Promotion Program, MEXT, Japan.
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