Open Journal of Organic Polymer Materials, 2012, 2, 45-52 Published Online July 2012 (
Stimulation of Selected Cellulases from Trichoderma reesei
with White Linearly Polarized Light
Ewelina Nowak1, Gohar Khachatryan1, Agnieszka Polit2, Lidia Krzeminska-Fiedorowicz1,
Marta Dziedzicka-Wasylewska2, Maciej Fiedorowicz1*
1Department of Chemistry and Physics, University of Agriculture, Krakow, Poland
2Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, Krakow, Poland
Email: *
Received April 17, 2012; revised May 23, 2012; accepted June 4, 2012
The impact of the illumination with white linearly polarized light (WLPL) of two commercially available cellulases
from Trichoderma reesei on their activity in hydrolysis of microcrystalline cellulose was studied. Enzymes were illu-
minated with WLPL for 60 min and 120 min and for each native and illuminated enzyme sample specific activity and
kinetics of enzyme catalyzed hydrolysis of microcrystalline cellulose were established. Molecular weight Mw and radii
if gyration Rg of protein chains of native and illuminated enzymes were measured by means of high pressure size exclu-
sion chromatography coupled with multiangle laser light scattering and refractometric detectors (HPSEC-MALLS-RI).
Conformations of protein chains of native and illuminated enzymes were evaluated on the basis of their circular dichro-
ism (CD) spectra. Additionally, molecular weight Mw and radii of gyration Rg of polysaccharide chains of microcrystal-
line cellulose native and digested for 10 min, 480 min and 1440 min with original and WLPL stimulated enzymes WT
and TR were taken. Illumination with WLPL of both cellulases studied did not change secondary structures of protein
molecules of native enzyme. Molecular weight Mw and radii of gyration Rg of illuminated enzymes differed greatly
from those found for native enzymes. Illumination of enzymes led to increase of specific activity and rate constants of
reaction of hydrolysis microcrystalline cellulose catalyzed by illuminated enzymes as compared with native enzymes.
Keywords: Cellulases; Polarized Light; Microcrystalline Cellulose
1. Introduction
Cellulose is the most abundant and renewable resource of
the biosphere. Degradation of cellulose, particularly that
enzymatic, to soluble sugars for fuels and industrially
essential chemicals has been expected to solve problems
of the energy shortage and, at the same time, environ-
ment protection. Thus far, combustion of wood is a prin-
cipal way of the cellulose utilization.
Low specific activity and high cost of polysaccharide
hydrolyzing enzymes [1-3] are the main limiting factors
in industrial hydrolysis of biomass containing cellulose.
Therefore, attempting to introduce efficient enzymes is
well rationalized. Hydrolysis of crystalline cellulose is
the rate-limiting step in the biomass conversion to etha-
nol because aqueous enzyme solutions cannot access the
contact with active centers of this insoluble, highly or-
dered structure. In the rate-limiting step disruption of a
single substrate chain from its native matrix takes place
instead of catalytic cleavage of the hydrolysis of the ma-
terial, thereby rendering it accessible to the catalytically
active cellulase site. Understanding cellulase structure
function relationships [4] is one of the key problems of
successful implementation of novel enzymes. Similarly,
understanding of microcrystalline cellulose molecular
structure and interactions between polysaccharide chain
with enzyme proteins will allow tailoring enzymes of
improved hydrolytic properties.
In the last 20 years, white linearly polarized light
(WLPL) has found many applications in medicine.
Medical cases being treated with polarized light include
severe second degree burns [5,6], wound healing [7], leg
ulcers, psoriasis and egzema [8,9]. The biostimulating
action of low-energy laser light, which is widely used in
medicine, could be attributed to the polarization of the
laser light [10]. Recently a stimulating influence of white
linearly polarized light (WLPL) on enzymes belonging to
hydrolyases family i.e. α-amylase in starch α-amylolysis
[11], xylanase in degradation of xylan [12], chitinase and
chitosanase in degradation of chitin and chitosan [13]
was reported. It was shown that WLPL action also
stimulates catalytic activity of cyclodextrin glucosyl-
transferase [14] and glucose oxidase [15] belonging to
*Corresponding author.
opyright © 2012 SciRes. OJOPM
cyclase and oxidase families respectively. The mecha-
nisms of the enzyme activity enhancement with WLPL is
unknown. It has been demonstrated [11] that 1 hr illumi-
nation with WLPL of α-amylase solutions led to statisti-
cally significant changes in the protein conformation of
illuminated enzyme, accompanied by acceleration of the
potato starch hydrolysis catalyzed by the activated en-
zyme. Such results indicate that the enhancement of the
activity of WLPL stimulated enzymes could be related to
conformational changes of the protein molecules.
In this study this problem is approached by examina-
tion of the effect of illumination with WLPL of two
commercially available cellulases, isolated from different
microorganisms. This effect is checked involving kinet-
ics of the hydrolysis of microcrystalline cellulose cata-
lysed with so stimulated enzymes. Absolute molecular
weight and conformation of protein chains of native and
illuminated enzymes was also compared. Additionally,
changes of molecular weight of cellulose chains in the
course of enzymatic degradation were recorded, to check
the enzyme source and/or WLPL stimulation have any
impact upon the molecular structure of enzymatically
degraded polysaccharide chains.
2. Experimental
2.1. Materials
Cellulase (TR) isolated from Trichoderma reesei (EC in aqueous solution (TR) and as a solid powder
(WT) was purchased from Sigma (Poznań, Poland) and
Worthington Biochemical Corporation (Lakewood NJ,
USA), respectively.
Microcrystalline cellulose and N,N-dimethylacetamide
(DMAc, HPLC grade) was purchased from Sigma, Poz-
nan, Poland.
2.2. Stimulation of the Enzymes by WLPL
Cellulase WT batch solution was prepared by dissolving
20 mg solid enzyme in 20 ml acetate buffer (pH = 5). In
the case of cellulase TR 2 ml aliquot of original enzyme
preparation was dissolved in 18 ml pH 5 acetate buffer.
Solutions of both enzymes were divided into three 6 ml
samples. One sample of each enzyme solution was stored
in the dark, whereas the other samples were placed in the
glass cells and illuminated with WLPL for 60 and 120
min respectively. Samples were illuminated from the 30
cm distance with a KB 502 slit illuminator (Kabid,
Chorzów, Poland) equipped with 150 W xenon arc (XBO
150. Oriel, Maidstone, UK). An HN 22 linear polarizing
filter (Polaroid, Waltham MA, USA) with a glass filter
cutting off wavelengths below 500 nm was mounted be-
tween the slit illuminator and the sample. The light
source emitted continuous radiation in the visible range.
Its energy flux at the position of the sample was 8
mW/c m2 as checked by YSI radiometer (Yellow Springs,
OH, USA). During the illumination temperature in the
place of the samples was controlled and did not exceeded
20˚C. Immediately after termination of the illumination
specific activity of each enzyme sample was measured.
2.3. Determination of Specific Activity of Native
and WLPL Stimulated Enzymes
Specific activity of enzymes prior to and after WLPL
illumination was determined by its effect on microcrys-
talline cellulose with respect to glucose formation. Re-
leased glucose was determined in a hexokinase/glucose-6-
phosphate dehydrogenase system at 340 nm using Glu-
cose (HK) Assay Kit (SIGMA, Poznan, Poland) em-
ploying procedure reported elsewhere [16]. One unit of
activity releases 0.01 mg glucose per hour from micro-
crystalline cellulose at 37˚C.
2.4. Enzymatic Hydrolysis of Microcrystalline
Cellulose Catalyzed by Native and
Illuminated Enzymes
Cellulose samples (800 mg) were exactly weighed in
Erlenmeyer flasks and suspended in acetate buffer (pH =
5, 38 ml). Erlenmeyer flasks containing cellulose sus-
pensions were placed in water bath for 5 min at 37˚C,
than aliquots (2 ml) of native and WLPL illuminated
solutions of respective enzymes were added and reaction
mixtures were incubated in a shaking water bath. In the
specified time intervals 0.4 ml aliquots were removed
from the reaction mixture, centrifuged at 8000 rpm for 10
min and supernatants were diluted with deionized water
to obtain final sample volume (2 ml). In each sample
reducing sugars were determined by the DNS method
2.5. Determination of Molecular Weight (Mw)
and Radii of Gyration (Rg) of, Native and
Enzymatically Digested Microcrystalline
Cellulose Chains
2.5.1. Sample Preparation for Chromatographic
Microcrystalline cellulose samples were dissolved in 8%
LiCl/DMac by slightly modified method described else-
where [17,18]. Thus, 4 × 10–2 g cellulose was placed in
10 ml measuring flask then cellulose was activated by two
consecutive swellings in 10 ml deionised water at 40˚C,
for 8 hr, followed by two consecutive transfers of the sam-
ples into 8 ml methanol for 45 min and finally into two
consecutive transfers into 8 ml DMAc for 12 hr. After
each exchange, solvent was removed by vacuum filtra-
Copyright © 2012 SciRes. OJOPM
tion. After the last DMAc exchange 10 ml of 8% LiCl/
DMAc was added to the sample. The samples were stirred
initially at room temperature for 24 hr, than stirring con-
tinued at 4˚C up to complete dissolution (2 to 3 days). Af-
ter dissolution 2 ml aliquots were taken for chroma-
tographic analysis, diluted to 20 ml volume with LiCl/
DMAc and filtered thorough a 0.5 μm pore PTFE filter
(Whatman, England) prior to the injection.
2.5.2. Enzyme Digested Cellulose Samples
Aliquots (2 ml) of the reaction mixture were collected at
time intervals (10 min, 8 hr, 24 hr), placed in boiled water
bath for 5 min, than centrifuged from denatured enzyme
protein. Supernatant was slowly evaporated to dryness at
60˚C and resulted solid was dissolved in the same manner
as native microcrystalline cellulose, omitting step of the
cellulose activation by solvent exchange with water and
Before analysis each sample was diluted to 20 ml vol-
ume with LiCl/DMAc and filtered thorough a 0.5 μm pore
PTFE filter (Whatman, England).
Anhydrous DMAc, carefully dried over aluminium so-
dium silicate molecular sieve, 0.4 nm effective pore size
(Fisher Scientific, Springfield, NJ, USA), was used for
activation, and for the preparation of the dissolution sol-
vent and SEC mobile phase. LiCl was oven dried and,
prior to use, stored in desiccator in pre-weighted amounts.
2.5.3. High Performance Size Exclusion
Chromatography (HPSEC-MALLS-RI)
The high performance size exclusion chromatography
(HPSEC) system for determination of average molecular
weight and radii of gyration of native and digested
microcrystalline cellulose chains consisted of a pump
(Shimadzu 10AC, Tokyo, Japan), an injection valve with
500 μl loop (model 7021, Rheodyne, Palo Alto, CA,
USA), a SDV guard column (50 × 8 mm, 20 μm, PSS,
Mainz, Germany), and two connected size exclusion
columns SDV (300 × 8mm, 20 μm, PSS, Mainz, Ger-
many) and SDV Linear XL (600 × 8 mm, 10 μm, PSS,
Mainz, Germany). The columns were placed in a ther-
mostated column compartment and the system was oper-
ated at 60˚C. A multiangle laser light scattering detector
(MALLS) operating in chromatographic mode using a
He-Ne laser light source (630.0 nm) (Dawn-DSP-F,
Wyatt Technology, Santa Barbara, CA, USA) and a dif-
ferential refractive index detector (L-7490, Merck, Dar-
mstadt, Germany) were connected to the columns. The
mobile phase (0.5% LiCl/DMAc) was filtered through 0.2
μm PTFE (Whatman, England) and vacuum degassed.
The flow rate of the mobile phase and the sample injec-
tion volume were 1.0 ml/min and 500 μl, respectively.
The output voltage of refractive index (RI) and light
scattering (LS) at 18 angles was used for calculation of
the weight-average molecular weight (Mw) and radius of
gyration (Rg) using Astra 4.73.04 software (Wyatt Tech-
nology, Santa Barbara, CA, USA). In calculations of Mw
and Rg dn/dc value 0.077 (ml/g) for cellulose [19] was
The Berry plot with third order polynomial fit was ap-
plied in the calculation of Mw and Rg values. Separations
were run in duplicates.
2.6. Determination of Molecular Weight (Mw)
and Radii of Gyration (Rg) of Enzyme
Protein Chains.
Average molecular weight (Mw) and radii of gyration (Rg)
of original and illuminated enzyme protein chains were
determined using HPSEC-MALLS-RI system described
in previous section.
The separation was carried out on a set of two con-
nected size exclusion columns TSKgel GMPWXL (300
7.8 mm, Tosoh Corporation, Tokyo, Japan) and TSKgel
2500 PWXL (300 7.8 mm, Tosoh Corporation, Tokyo,
Japan). The columns and the refractive index detector
were maintained at 25˚C. The mobile phase (0.15 M
NaNO3 with 0.02% sodium azide) was filtered through
0.2 and 0.1 μm cellulose acetate filters (Whatman, Eng-
land) and vacuum degassed. The flow rate of the mobile
phase and the sample injection volume were 0.4 ml/min1
and 500 μl respectively. Prior to injection, the batch solu-
tions of original and illuminated enzymes were filtered
through a 0.5 μm pore cellulose acetate filter (Whatman,
2.7. Evaluation of Protein Conformation of
Enzymes Cd Spectra
Circular dichroism spectra of the WT and TR cellulase
solutions kept in the dark and illuminated for 1 h and 2 h
with visible nonpolarized and linearly polarized light
were recorded using a JASCO J-710 (Jasco, Japan) spec-
trophotometer (upgraded to a J-715).
Sample Preparation
Water batch solutions (30 ml) of each enzyme at concen-
tration of 2 mg/ml were divided into 3 parts. One sample
was stored in the dark. Two other samples of each en-
zyme were illuminated with WLPL for 1 h and 2 h, re-
spectively, under conditions described above. Immedi-
ately after termination of illuminations samples were
dialyzed and protein concentration in each sample was
determined. Samples were diluted with water to obtain
final protein concentration of 0.7 mg/ml. For evaluation
of the secondary protein structure, CD spectra of each
enzyme sample were recorded in the range of 250 - 190
nm using quartz cylindrical cells with the l of 0.2 mm
optical path and scanning speed 20 nm/min. Subse-
Copyright © 2012 SciRes. OJOPM
quently, in order to evaluate the tertiary protein structure,
spectra were recorded in the range of 350 - 250 nm using
a quartz cylindrical 1 cm optical path cell. Each spectrum
was the average of three scans with the average buffer
control spectrum subtracted. Data obtained were not
smoothed. Proteins secondary structure content was per-
formed with 1.51 Spectra Manager software using refer-
ence set containing five proteins: hemoglobin, lysozyme,
myoglobin, ribonuclease A and α-chymotripsin A.
3. Results and Discussion
Specific activity of WT and TR enzymes, original and
WLPL illuminated for 60 (WT60, TR60) and 120 min
(WT120, TR120) are presented in Table 1.
Specific activity of enzyme WT illuminated with
WLPL for 60 min was slightly, although statistically sig-
nificantly, higher, than activity of original enzyme. Pro-
longed (120 min) illumination of this sample led to fur-
ther, more pronounced, increase in the enzyme specific
activity. In the case of enzyme TR, Illumination with
WLPL for 60 min did not change specific activity of TR
enzyme. However, its illumination for 120 min led to
significant increase in its specific activity.
Hydrolysis kinetics curves of microcrystalline cellu-
lose digested by solutions of non-illuminated and WLPL
illuminated enzymes WT and TR are presented in Fig-
ures 1(a) and (b) respectively.
Enzymes degrading carbohydrate polymers can follow
three fundamentally different mechanisms [20]. They are
1) a multiple chain mechanism, where the enzyme-sub-
strate complex dissociates after each reaction; 2) a sin-
gle-chain mechanism, where the enzyme remains associ-
ated with the substrate until every cleavage-plaint linkage
in the chain hydrolyzes; and 3) a multiple attack mecha-
nism, where a given average number of attacks were per-
formed after the formation of the enzyme-substrate com-
plex. As it can be seen in Figures 1(a) and (b), the deg-
radation of cellulose by original and illuminated enzymes
follows two rate laws.
Rate constants for both stages of enzymatic degrada-
tion of cellulose with native and WLPL stimulated en-
zymes WT and TR are given in Table 2 Illumination of
enzyme WT with WLPL for 60 min led to slight, al
though statistically significant rise of the rate constant for
the first stage of cellulose degradation. Rate constant for
the first step of the cellulose hydrolysis catalyzed by WT
illuminated for 120 min remained unchanged. In the case
of enzyme TR, illumination with WLPL for 120 min led
to significant rise of rate constant of the first stage of the
cellulose degradation.
In the case of the second, much slower stage of the cel-
lulose degradation, illumination of both WT and TR en-
zymes with WLPL for 120 min led to a rise of k2 rate con-
stant. Acceleration of the second step of the cellulose deg-
radation was much more pronounced in the case of en-
zyme WT stimulated with WLPL for 120 min as com-
pared to similarly treated enzyme TR.
In order to check whether stimulation of cellulases by
means of illumination with WLPL have any impact on
molecular structure of cellulose chains in the course of
hydrolysis average molecular weight (Mw) and radii of
gyration (Rg) of microcrystalline cellulose chains native
and digested for 10min, 480 min and 1440 min with
original and WLPL stimulated enzymes WT and TR were
taken. Measurements were performed for the material
eluted under whole polysaccharide peak from HPLC col-
umn (Table 3).
Table 1. Specific activity of enzymes WT and TR prior to
and after illumination with the WLPLa.
Enzyme specific activity
Enzyme illumination time
[min] WT [u/mg] TR [u/ml]
0 65.0 ± 0.6 820.1 ± 10.0
60 68.3 ± 0.8 823.0 ± 9.1
120 73.4 ± 0.9 1166.6 ± 10.1
aMeans of three independent experiments ± standard deviation.
Figure 1. (a) Kinetics of the microcrystalline cellulose hy-
drolysis with enzyme WT; (b) Kinetics of the microcrystal-
line cellulose hydrolysis with enzyme TR. Samples of en-
zymes native () and illuminated with WLPL for 60 min ()
and 120 min ().
Copyright © 2012 SciRes. OJOPM
Copyright © 2012 SciRes. OJOPM
Table 2. Rate constants of the enzymatic degradation of cel-
lulose with enzymes WT and TR prior to and after their
stimulation with WLPLa.
Samplea k1 × 104[mg/mL–1/ min–1] k2 × 105[mg/mL–1/min–1]
WT 2.60 ± 0.20 4.34 ± 0.21
WT 60 3.00 ± 0.15 4.34 ± 0.32
WT 120 2.60 ± 0.15 8.68 ± 0.34
TR 7.81 ± 0.20 1.30 ± 0.13
TR 60 7.38 ± 0.20 1.30 ± 0.12
TR 120 9.12 ± 0.18 1.74 ± 0.21
aMeans of three independent experiments ± standard deviation.
Molecular weight of polysaccharides digested for 10
min by original enzymes WT and TR slightly declined.
The same effect was noted for the cellulose molecules
digested for 10 min by WT and TR enzymes stimulated
with WLPL for 60 min. However, average molecular
weight of cellulose chains digested 10 min by WT and TR
illuminated with WLPL for 120 min decreased signifi-
cantly to 1.58 × 106 and 1.57 × 106, respectively. Calcu-
lated average molecular weight of cellulose chains left
non-digested in the reaction mixture after 480 min of the
hydrolysis differed greatly, depending of the enzyme ori-
gin and time of their stimulation with WLPL. Sharp reduc-
tion of average molecular weight of the cellulose polysac-
charide chains digested for 480 min by original WT (0.761
× 106), WT60 (0.550 × 106) and WT120 min (0.148 × 106)
could be noted. Likely, original and WLPL stimulated
enzyme in the first stage of the reaction, hydrolyzed
mainly long cellulose chains of high molecular weight. In
contrast to it, average molecular weight of polysaccharide
chains digested for 480 min by original enzyme TR (2.56
× 106) was slightly lower than molecular weight observed
for cellulose molecules digested by this enzyme for 10 min
(2.88 × 106). Possibly, mainly shorter cellulose chains
were attacked by original enzyme TR in the first stage of
hydrolysis. Similarly, such assumption could explain that
in the reaction catalyzed by TR 120 (1.99 × 106) average
molecular weight of non-digested cellulose molecules was
higher in the reaction mixture hydrolyzed for 480 min than
in the reaction mixture digested for 10 min. Interestingly,
the 60 min stimulation with WLPL led to significant de-
crease in the average molecular weight of cellulose chains
left in the reaction mixture after 480 min digestion by
TR60 (0.330 × 106). Average molecular weight of cellu-
lose chains remaining in the reaction mixture after com-
pleting hydrolysis within 1440 min were much higher than
molecular weight of polysaccharide chains left after 480
min digestion for all original and WLPL enzymes studied.
It could indicate that in the second stage of hydrolysis en-
zymes digested mainly short cellulose chains. Molecular
weight of original and WLPL stimulated enzymes WT and
TR are presented in Table 4 and results of evaluation of
the enzymes protein secondary conformation are summa-
rized in Table 5.
Table 3. Average molecular weight Mw and radii of gyration Rg of the cellulose chains hydrolyzed by native and illuminated
with WLPL enzymes WT and TR prior to and after stimulation with WLPL.a
Average molecular weight (Mw) and radii of gyration (Rg) of enzyme digested cellulose chains
Hydrolysis time
0 10 480 1440
MW × 106 R
g [nm] MW × 106 R
g [nm] MW × 106 R
g [nm] MW × 106 R
g [nm]
WT native 2.99 ± 0.104 80.15 ± 5.15 2.65 ± 0.105 59.7 ± 4.5 0.761 ± 0.05042.7 ±3.1 0.855 ± 0.025 93.1 ± 3.1
WT 60 2.99 ± 0.104 80.1 ± 5.15 2.67 ± 0.099 40.0 ± 5.0 0.550 ± 0.03447.7 ±3.4 1.35 ± 0.034 59.1 ± 5.0
WT 120 2.99 ± 0.104 80.1± 5.15 1.58 ± 0.099 103.8 ± 5.10.148 ± 0.04054.4 ±2.5 1.99 ± 0.099 39.8 ± 4.5
TR native 2.99 ± 0.104 80.1± 5.15 2.88 ± 0.100 78.3 ± 5.5 2.56 ± 0.09956.3 ± 3.3 1.61 ± 0.87 50.8 ± 4.1
TR 60 2.99 ± 0.104 80.1 ± 5.15 2.19 ± 0.102 42.1 ± 4.1 0.355± 0.02047.8 ± 2.6 2.45 ± 0.099 50.2 ± 2.1
TR 120 2.99 ± 0.104 80.1 ± 5.15 1.57 ± 0.080 40.6 ± 3.9 1.99 ± 0.05854.4 ± 3.2 2.52 ± 0.105 52.8 ± 2.3
aMeans of three independent experiments ± standard deviation; bThe numbers denotes illumination time (min).
Table 4. Molecular weight of native and WLPL illuminated
enzymes WT and TR.a
Enzyme molecular weight
Enzyme illumination
time [min] WT Mw × 104 TR Mw × 104
0 3.62 ± 0.25 4.04 ± 0.22
60 5.54 ±.0.31 6.25 ± 0.30
120 3.82 ± 0.29 4.67 ± 0.29
aMean of three independent experiments ± standard deviation.
Table 5. Evaluation of the secondary structure of protein
chains of enzymes WT and TR prior to and after WLPL il-
lumination for 60 and 120 min.
Structure WT WT60 WT120 TR TR60TR120
Helix 18.6 18.7 18.0 29.5 27.4 29.6
Beta 34.5 34.3 34.7 29.3 30.5 29.2
Turn 5.8 6.2 7.1 0.0 0.0 0.0
Random 41.0 40.8 40.2 41.2 42.1 41.2
aThe numbers denotes illumination time.
In this work we decided to measure molecular weight
of original and illuminated enzymes and, based on re-
corded CD spectra of the enzymes, evaluate their secon-
dary conformation. Illumination with WLPL signifi-
cantly changed molecular weight of the enzyme protein
molecules. Molecular weight of original WT (3.63 × 104)
and TR (4.04 × 104) enzymes increased significantly af-
ter 60 min illumination to 5.54 × 104 and 6.25 × 104 re-
spectively. Further, 120 min illumination of the enzyme
solutions led to decrease in the molecular weight of both
WT120 (3.82 × 104) and TR120 (4.67 × 104) enzymes.
Using Astra 4.73.04 software both, RI and LS detectors
provided differential molar mass distribution. The dif-
ferential molar mass distribution offered the amount of
polymer (differential weight fraction) in a given molar
mass interval [21]. Plots of differential weight fraction vs
molar mass for original and illuminated WT and TR en-
zymes are given in Figures 2(a) and (b), respectively.
Clearly, illumination with WLPL changed the distribu-
tion of molecular weight of the enzyme WT protein
Polydispersity of molecular weight of the original en-
zyme protein chains was much smaller than polydisper-
sity observed for enzyme samples illuminated for 60 and
120 min. Samples of illuminated enzymes contained frac-
tions with protein chains with much lower and much
higher molecular weight than these in original enzymes.
It has been established that the cellulase complex of
Trichoderma reesei contains at least three enzyme com-
ponents that are both physically and enzymatically dis-
tinct and that all three components play essential roles in
the overall process of converting cellulose to glucose.
Figure 2. (a) Plot of Differential Weight Fraction vs Molar
Mass for enzyme WT proteins prior to (WT native), and
after illumination with WLPL for 60min (WT60) and 120
min (WT120); (b) Plot of Differential Weight Fraction vs
Molar Mass for enzyme TR proteins prior to (TR native)
and after illumination with WLPL for 60 min (TR60) and
120 min (TR 120).
Observed increase of polydispersity of molecular weight
of protein chains in WLPL illuminated samples of en-
zyme WT could be result of light induced association
between protein chains belonging to three different
components of enzyme complex. One could expect that
self association of protein molecules should lead to the
products with molecular weight values periodic as multi-
ples of a base molecular weight. However, random asso-
ciation, induced by WLPL, between three different pro-
tein chains led to the association products exhibiting
broad range of molecular weight.
Data presented in Table 5 revealed that independently
the illumination time the secondary structure of activated
proteins in cellulases WT and TR did not change.
It has been shown [11] that pre-illumination of α-amylase
Copyright © 2012 SciRes. OJOPM
solutions with WLPL enhanced the starch hydrolysis rate
and extent. Simultaneously, the illumination of α-amy-
lase led to significant changes in the secondary confor-
mation of the enzyme protein molecule. It has been as-
sumed that conformational changes in the enzyme pro-
tein structure were, at least partially, responsible for the
observed enhancement of hydrolytic activity of pre-il-
luminated α-amylase. Protein compositions of α-amylase
and cellulases WT and TR are entirely different. Amy-
lase is a single chain glycoprotein of about 475 residues
and molecular weight in the range 50,000 [22]. Cellu-
lases WT and TR are complex of three physically distinct
components [23]. Such differences in molecular structure
between amylase and cellulases studied could explain
lack of conformational changes in proteins of WLPL
activated cellulases.
4. Conclusions
1) Stimulation with white linearly polarized light of
two cellulases WT and TR isolated from Trichoderma
reesei increases the enzyme specific activity of the
microcrystalline cellulose digestion and rate constants of
both stages of enzymatic hydrolysis of microcrystalline
2) The WLPL stimulation of cellulases significantly
influences molecular structure of cellulose chains in di-
gested microcrystalline cellulose compared to these re-
sulting from the hydrolysis with original enzyme.
3) Illumination of both WT and TR enzymes led to
significant changes of molecular weight of enzyme protein
chains, however, activation of enzymes with WLPL did
not change enzyme protein secondary structure.
5. Acknowledgements
This work was financed by grant N N313 265936 from
National Science Center, Poland.
[1] K. Wickholm, E. Hult, P. Larsson, T. Iversen and H.
Lennholm, “Quantication of Cellulose Forms in Com-
plex Cellulose Materials: A Chemometric Model,” Cel-
lulose, Vol. 8, No. 2, 2001, pp. 139-148.
[2] L. R. Lynd, “Overview and Evaluation of Fuel Ethanol
from Cellulosic Biomass: Technology, Economics, the
Environment, and Policy,” Annual Review of Energy and
the Environment, Vol. 21, 2001, pp. 403-465.
[3] L. R. Lynd, R. T. Elander and C. E. Wyman, “Likely
Features and Costs of Mature Biomass Ethanol Technol-
ogy,” Applied Biochemistry and Biotechnology, Vol. 57-
58, 1996, pp. 741-761. doi:10.1007/BF02941755
[4] L. Laureano-Perez, F. Teymouri, H. Alizadeh and B. E.
Dale, “Understanding Factors That Limit Enzymatic Hy-
drolysis of Biomass,” Applied Biochemistry and Bio-
technology, Vol. 121, 1996, pp. 1081-1100.
[5] H. Hoeksema, S. Monstrey, K. Van Landuyt, Ph. Blon-
deel, P. Tonnard and A. Verpaele, “The Use of Polarised
Light in the Treatment of Severely Burned Patients (Ab-
stract),” 10th Congress of the International Society for
Burn Injuries, Jerusalem, 1-6 November 1998, pp. 1-6.
[6] K. Depuydt, S. Monstrey and H. Hoeksma, “The Stimu-
lating Effects of Polarized Light on Wound Healing and
Avoiding Surgery in the Treatment of Deep Dermal Burn
Wounds Using Polarized Light,” 10th Annual Meeting of
the European Association of Plastic Surgeons, Madrid, 21
May 1999, pp. 21-25.
[7] W. Vanscheidt, “The Effect of Polarized Light on Wound
Healing,” European Journal of Plastic Surgery, Vol. 24,
No. 8, 2002, pp. 383-390.
[8] E. Bazso, Sz. Varju, P. Szego, K. Roza and P. Apai, “Ap-
plication of Incoherent Wide Band Polarised Light to
Promote Healing of Wounds,” Central Research Institute
for Physics, Budapest, 1982, pp. 121-130.
[9] W. Stegmann, “Behandlung des Ulcus Cruris mit Polar-
isiertem Licht,” Phlebologie und Proktologie, Vol. 14,
1985, pp. 96-97.
[10] M. Fenyö, “Theoretical and Experimental Basis of Bio-
stimulation,” Optics & Laser Technology, Vol. 16, 1984,
pp. 209-215. doi:10.1016/0030-3992(84)90029-X
[11] M. Fiedorowicz and G. Chaczatrian, “Effect of Illumina-
tion with the Visible Polarized and Non-Polarized Light
on α-Amylolysis of Starches of Different Botanical Ori-
gin,” Journal of Agricultural and Food Chemistry, Vol.
51, No. 26, 2003, pp.7815-7819. doi:10.1021/jf026202r
[12] A. Konieczna-Molenda, V. M. F. Lai, M. Fiedorowicz, G.
Khachatryan and P. Tomasik, “Effect of Linearly Polar-
ized Light upon Xylanase Activity,” Biotechnology Pro-
gress, Vol. 24, No. 2, 2008, pp. 385-388.
[13] A. Konieczna-Molenda, M. Fiedorowicz, W. Zhong and P.
Tomasik, “Polarized Light-Stimulated Enzymatic Hy-
drolysis of Chitin and Chitosan,” Carbohydrate Research,
Vol. 343, No. 18, 2008, pp. 3117-3119.
[14] M. Fiedorowicz, A. Konieczna-Molenda and G. Khacha-
tryan, “Stimulation of Cyclodextrin-Glycosyltransferase
(Turozyme) Activity by Illumination with Linearly Po-
larized Visible Light,” Biotechnology Progress, Vol. 25,
No. 1, 2009, pp. 147-150. doi:10.1002/btpr.90
[15] A. Konieczna-Molenda, M. Fiedorowicz and P. J. Toma-
sik, “Stimulation of Glucose Oxidase with White Linearly
Polarized Light,” Biotechnology Progress, Vol. 26, No. 2,
2010, pp. 393-396.
[16] G. L. Miller, “Use of Dinitrosalicylic Acid Reagent for
Determination of Reducing Sugars,” Analytical Chemis-
try, Vol. 31, No. 3, 1959, pp. 426-428.
[17] A. Dupont and G. Harrison, “Conformation and dn/dc
Determination of Cellulose in N,N-Dimethylacetamide
Copyright © 2012 SciRes. OJOPM
Copyright © 2012 SciRes. OJOPM
Containing Lithium Chloride,” Carbohydrate Polymers,
Vol. 58, 2004, pp. 233-243.
[18] A. Dupont and G. Harrison, “Comparative Evaluation of
Size-Exclusion Chromatography and Viscometry for the
Characterisation of Cellulose,” Journal of Chromatogra-
phy A, Vol. 1026, No. 1, 2004, pp. 129-141.
[19] “Worthington Enzyme Manual,” Worthington Biochemi-
cal Corporation, Lakewood, 1993.
[20] D. W. Schrott, “Differential Molecular Weight Distribu-
tions in High Performance Size Exclusion Chromatogra-
phy” Journal of Liquid Chromatography & Related Te-
chnologies, Vol. 16, No. 16, 1993, pp. 3371-3391.
[21] S. J. Horn, A. Sørbotten, B. Synstad, P. Sikorski, M. Sør-
lie, K. M. Vårum and V. G. H. Eijsink, “Endo/Exo
Mechanism and Processivity of Family 18 Chitinases Pro-
duced by Serratia marcescens,” FEBS Journal, Vol. 273,
No. 3, 2006, pp. 491-503.
[22] M. Granger, B. Abadie and G. Marchis-Mouren, “Limited
Action of Trypsin on Porcine Pancreatic Amylase: Char-
acterization of the Fragments,” FEBS Letters, Vol. 56, No.
2, 1975, pp. 189-197.
[23] K. King and M. Vessal, “Enzymes of the Cellulase Com-
plex,” Advances in Chemistry, Vol. 95, 1969, pp. 130-