Improved peripheral nerve regeneration in streptozotocin-induced
diabetic rats by oral lumbrokinase
Han-Chung Lee1,2, Yuan-Man Hsu3, Chin-Chuan Tsai4,5, Cherng-Jyh Ke6,
Chun-Hsu Yao7,8, Yueh-Sheng Chen7,8†
1
Graduate Institute of Clinical Medical Science, China Medical University, Taiwan
Division of Neurosurgery, China Medical University Hospital, Taichung, Taiwan
3
Department of Biological Science and Technology, China Medical University,
Taichung, Taiwan
4
School of Chinese Medicine for Post-Baccalaureate, I-Shou University,
2
Kaohsiung, Taiwan
5
Chinese Medicine Department, E-DA Hospital, Kaohsiung, Taiwan
6
Department of Orthopedics, College of Medicine, National Taiwan University, Taipei,
Taiwan
7
School of Chinese Medicine, China Medical University, Taichung, Taiwan
8
Department of Biomedical Informatics, Asia University, Wufeng District, Taichung,
Taiwan
Running Title: Improved nerve regeneration in diabetic rats by oral lumbrokinase
*Corresponding author: Yueh-Sheng Chen, PhD
School of Chinese Medicine, China Medical University,
#91, Hseuth-Shih Road, Taichung 40402, Taiwan
Tel.: 886-4-22053366 ext. 3308; Fax: 886-4-22032295
E-mail: yuehsc@mail.cmu.edu.tw
Abstract
We assessed the therapeutic effects of lumbrokinase, a group of enzymes extracted
from the earthworm, on peripheral nerve regeneration using well-defined sciatic nerve
lesion paradigms in diabetic rats induced by injection of streptozotocin. We found that
lumbrokinase therapy could improve the rats’ circulatory blood flow and promote the
regeneration of axons in a silicone rubber conduit after nerve transection.
Lumbrokinase treatment could also improve the neuromuscular functions with better
nerve conductive performances. Immunohistochemical staining showed that
lumbrokinase could dramatically promote calcitonin gene-related peptide expression
in lamina I-II regions in the dorsal horn ipsilateral to the injury and cause a marked
increase in the number of macrophages recruited within the distal nerve stumps. In
addition, the lumbrokinase could stimulate the secretion of interleukin-1, nerve
growth factor, platelet-derived growth factor, and transforming growth factor-β in
dissected diabetic sciatic nerve segments. In conclusion, administration of
lumbrokinase after nerve repair surgery in diabetic rats was found to have remarkable
effects on promoting peripheral nerve regeneration and functional recovery.
KEYWORDS: lumbrokinase; peripheral nerve regeneration; earthworm
Introduction
Diabetes mellitus is a metabolic disorder that may cause axonal atrophy, nerve
demyelination, and delayed Wallerian degeneration and subsequent regeneration of
nerve fibers (Zochodne et al., 2007). Mechanisms that may account for diabetic nerve
regenerative failure include deficits in support of neurotrophins and neuropeptides,
abnormalities in regenerative microenvironment with ischemia, and impairments in
invasion of macrophages for phagocytosing axon and myelin debris following nerve
injury (Yasuda et al., 2003).
Previous studies have shown that several neurotrophic factors tested in animal
models of diabetic neuropathy with variable success in peripheral nerve regeneration,
such as the nerve growth factor (Unger et al., 1998) and the insulin-like growth factor
(Zhuang et al., 1996). Studies have also shown that traditional Chinese medicine
(TCM) may affect peripheral nerve regeneration in diabetic animals by promoting
Schwann cell proliferation and increasing expression of multiple neurotrophic factors
(Piao and Liang, 2012). Our group has demonstrated that administration of crude
extracts of earthworm could promote the regeneration of injured rat sciatic nerve
(Chen et al., 2010a). Earthworm, as a TCM, has been used thousands of years in
China (Cooper and Balamurugan, 2010). Lumbrokinase, a group of enzymes extracted
from the earthworm, has been identified, which could dissolve fibrin clot by
converting plasminogen to plasmin (Cho et al., 2004). Clinical studies indicate that
orally-administered lumbrokinase could improve regional myocardial perfusion in
patients with stable angina (Kasim et al., 2009) and have no obvious side effects on
nervous, respiratory, or circulatory systems (Cooper et al., 2004). It also could reduce
the proteinuria and improve the glomerulosclerosis and tubulointerstitial fibrosis in
diabetic rats (Sun et al., 2013).
In the present study, a diabetic rat model was established using streptozotocin
(STZ) injection. Sciatic nerves in the diabetic rats were transected and the severed
nerve ends were sutured into a 10 mm long silicone rubber tube. Therapeutic effects
of lumbrokinase on peripheral nerve regeneration in diabetic rats were then evaluated
by determination of their cutaneous blood flow, electrophysiological nerve function,
expression of calcitonin gene-related peptide (CGRP) in spinal cord, macrophages
recruited in nervous tissues, and morphometric observations of regenerated nerve
cables in the bridging chamber. Finally, changes of mRNA levels of interleukin-1
(IL-1), nerve growth factor (NGF), platelet-derived growth factor (PDGF), and
transforming growth factor-β (TGF-β) of diabetic rat sciatic nerve segments by adding
conditioned media of lumbrokinase to elucidate mechanisms underlying its
therapeutic effects.
Materials and Methods
Induction of Diabetes
Prior to the beginning of the in vivo testing, the protocol was approved by the ethical
committee for animal experiments of the China Medical University, Taichung, Taiwan.
Diabetes was induced in adult male Sprague-Dawley rats (250-300 g, BioLasco Co,
Ltd, Taipei, Taiwan) by tail vein injection of a single 50 mg/kg dose of STZ (Sigma
Chemical Co, St Louis, MO). The STZ was solubilized in normal saline immediately
before injection. Seven days after STZ injection, serum glucose measurements were
determined on all animals with a glucose analyzer (Accu-Chek, Roche, Basel,
Switzerland). Animals with an initial blood glucose of 300 mg/dl or greater qualified
as diabetic.
Nerve surgery
All rats were anesthetized with an inhalational technique (AErrane; Baxter, Deerfield,
IL) as reported elsewhere in rats (Lin et al., 2014). The right sciatic nerves were
severed into proximal and distal segments. The proximal stump was then secured with
a single 9-0 nylon suture through the epineurium and the outer wall of a silicone
rubber chamber (1.47 mm inner diameter, 1.96 mm outer diameter; Helix Medical, Inc,
Carpinteria, CA). The distal stump was secured into the other end of the chamber.
Both the proximal and the distal stumps were secured to a depth of 1 mm into the
chamber, leaving a 10-mm gap between the stumps. The muscle and skin were closed.
All animals were housed in temperature (22°C) and humidity (45%) controlled rooms
with 12-hour light cycles. They had access to food and water ad libitum.
Drug treatment
The diabetic rats were randomly allocated into 4 experimental groups. Control rats in
group A received PBS only. The rats in groups B-D were treated with lumbrokinase
(Boluoke®, CRNA, Canada) at concentrations of 300, 600, and 1200 µg/Kg dissolved
in 0.01 M of PBS every other day for 4 weeks using intragastric gavage.
Cutaneous blood flow measurement
The blood flow measurement was performed in a quiet room and the ambient
temperature was controlled at 25°C and humidity at 50% by using an air conditioner.
Each rat was maintained at a light stage of anesthesia and placed on a stainless steel
tray. Cutaneous blood flow in the hindlimb footpad ipsilateral to the injury of the rat
was measured with a laser Doppler flowmetry device (wavelength, 780 nm; DRT4;
Moor Instruments Ltd., Millwey, Axminster, UK) at various time points: 0 d, 14 d,
and 28 d after the nerve repair.
Electrophysiological Techniques
Four weeks after nerve repair, all animals were re-anesthetized and the sciatic nerve
exposed. The nerve was given a supramaximal stimulus through a pair of needle
electrodes placed directly on the sciatic nerve trunk, 5-mm proximal to the transection
site. Latency, amplitude, and area of the evoked muscle action potentials (MAPs)
were recorded from the gastrocnemius muscle with microneedle electrodes linked to a
computer (Biopac Systems, Inc., Goleta, California). The latency was measured from
stimulus to the takeoff of the first negative deflection. The amplitude and the area
under the MAP curve from the baseline to the maximal negative peak were calculated.
The MAP was then used to calculate the nerve conductive velocity (NCV), which was
carried out by placing the recording electrodes in the gastrocnemius muscles and
stimulating the sciatic nerve proximally and distally to the silicone rubber conduit.
The NCV was then calculated by dividing the distance between the stimulating sites
by the difference in latency time.
Histological Techniques
Immediately after the recording of muscle action potential, all of the rats were
perfused transacrdially with 150 ml normal saline followed by 300 ml 4%
paraformaldehtde in 0.1 M phosphate buffer, pH 7.4. After perfusion, the L4 spinal
cord and the distal stump outside the nerve gap were quickly removed and post-fixed
in the same fixative for 3-4 h. Tissue samples were placed overnight in 30% sucrose
for cryoprotection at 4°C, followed by embedding in optimal cutting temperature
solution. Samples were the kept at -20°C until preparation of 18 μm sections was
performed using a cryostat, with samples placed upon poly-L-lysine-coated slide.
Immunohistochemistry of frozen sections was carried out using a two-step protocol
according to the manufacturer's instructions (Novolink Polymer Detection System,
Novocastra). Briefly, frozen sections were required endogenous peroxidase activity,
was blocked with incubation of the slides in 0.3% H2O2, and nonspecific binding sites
were blocked with Protein Block (RE7102; Novocastra). After serial incubation with
rabbit- anti-CGRP polyclonal antibody 1:1000 (calbiochem, Germany), Post Primary
Block (RE7111; Novocastra), and secondary antibody (Novolink Polymer RE7112),
the L4 spinal cord sections were developed in diaminobenzidine solution under a
microscope and counterstained with hematoxylin. Similar protocols were applied in
the sections from the distal stump except they were incubated with anti-rat CD68
1:100 (AbD Serotec, Kidlington, UK). Sciatic nerve sections were taken from the
middle regions of the regenerated nerve in the chamber. After the fixation, the nerve
tissue was post-fixed in 0.5% osmium tetroxide, dehydrated, and embedded in Spurr’s
resin. The tissue was then cut to 2-µm thickness by using a microtome (Leica EM
UC6, Leica Biosystems, Mount Waverley, Australia) with a diamond knife and stained
with toluidine blue.
Changes in mRNA levels of IL-1, NGF, PDGF, and TGF-β of rat sciatic nerve
segments conditioned by lumbrokinase
Sciatic nerve segments (3 cm) of adult diabetic Sprague-Dawley rats were cultured in
1 ml Dulbecco’s modified Eagle’s medium supplemented with 10% fetal calf serum.
After three days culture, 600 µg/kg of lumbrokinase based on the weight of nerve
segment was dissolved in medium and added. Three hours later, total RNAs were
extracted from nerve tissues with TRIzol reagent and the amount of RNA estimated
by spectrophotometry at 260 nm. For real-time (RT)-PCR analysis, two-step RT-PCR
was carried out using a high-capacity cDNA reverse transcription kit (Applied
Biosystems, USA), and a 16S rRNA gene PCR assay was used as a housekeeping
gene control assay. The reactions were performed in 20 µl (total volume) mixtures
containing primers at a concentration of 400 nM. The reaction conditions consisted of
2 min at 50°C, 10 min at 95°C, then 40 cycles of 15 s at 95°C, followed by 1 min at
60°C. Melting curve analysis was used to determine the PCR specificity and was
performed using 80 10-s cycles, with the first cycle at 60°C and the temperature
increasing by 0.5°C for each succeeding cycle. All reactions were carried out in
triplicate from three cultures. Each assay was run on an Applied Biosystems 7300
Real-Time PCR system. The threshold cycle (Ct) is defined as the fractional cycle
number at which the fluorescence passes the fixed threshold data, and was determined
using the default threshold settings. Relative quantification of mRNA expression was
calculated using the 2-ΔΔCt method (Applied Biosystems User Bulletin N°2 (P/N
4303859)). Data were presented as the relative expression of target mRNA,
normalized with respect to GAPDH mRNA and relative to a calibrator sample that
was collected at 0 min of infection. The primers used in this study are shown in Table
1.
Image Analysis
All tissue samples were observed under an optical microscope (Olympus IX70;
Olympus Optical Co, Ltd, Tokyo, Japan) with an image analyzer system (Image-Pro
Lite; Media Cybernetics, Silver Spring, MD). CGRP-immunoreactivity (IR) in dorsal
horn in the lumbar spinal cord was detected by immunohistochemistry as described
previously (Lee et al., 2013). The immuno-products were confirmed positive-labeled
if their density level was over five times background levels. Under a 400x
magnification, the ratio of area occupied by positive CGRP-IR in dorsal horn
ipsilateral to the injury following neurorrhaphy relative to the lumbar spinal cord was
measured. The number of neural components in each nerve section was also counted.
As counting the myelinated axons, at least 30 to 50 percent of the sciatic nerve section
area randomly selected from each nerve specimen at a magnification of 400x was
observed. The axon counts were extrapolated by using the area algorithm to estimate
the total number of axons for each nerve. Similarly, macrophages were counted in
each nerve section at the distal stump. The density of myelinated axons and the
macrophages was then obtained by dividing the myelinated axon counts and the
macrophage counts by the total nerve areas, respectively.
Statistical Analysis
For
the
statistical
analysis
of
immunohistochemical,
morphometric,
and
electrophysiological measurements of regenerated nerves, data were collected by the
same observer and expressed as mean ± standard deviation, and comparisons between
groups were made by the 1-way analysis of variance (SAS 8.02). The Tukey test was
then used as a post hoc test. Statistical significance was set at P < 0.05.
Results
The cutaneous blood flow in the ipsilateral hindpaw to the injury in response to
lumbrokinase treated in diabetic rats at different time points measured is shown in
Figure 1. Mean blood flows recorded immediately after the nerve repair in the
lumbrokinase-treated animals and the controls were about 110 perfusion units and no
significant differences were seen among these animals. After 14 days of nerve repair,
the three groups of lumbrokinase-treated animals had a blood flow appropriately 135
perfusion units, which was significantly larger than that (120 perfusion units) in the
controls (p <0.05). After 28 days of nerve repair, the lumbrokinase-treated rats at 600
and 1200 µg/Kg still kept their blood flows around 135 perfusion units, compared to
128 perfusion units in those at 300 µg/Kg. Even though, all of these
lumbrokinase-treated rats still had a significantly larger cutaneous blood flow than the
controls (p < 0.05).
Electrophysiological results are shown in Figure 2. It was noted that
lumbrokinase-treated diabetic animals at 600 and 1200 µg/Kg had a significantly
larger NCV as compared to the controls (p < 0.05). A significantly smaller latency was
observed in the lumbrokinase-treated animals at 1200 µg/Kg as compared to that in
the controls (p < 0.05). In addition, the lumbrokinase-treated animals at 600 µg/Kg
had significantly larger amplitudes and MAP areas versus those in the control group
(p < 0.05).
Immunohistochemical staining showed that CGRP-labeled fibers were seen in
the area of lamina I-II regions in the dorsal horn ipsilateral to the injury in all of the
rats (Figure 3). Compared to the controls, it was noted that the ratio of area occupied
by positive CGRP-IR was dramatically increased in the diabetic animals after
receiving the lumbrokinase treatment, especially at dosages of 600 and 1200 µg/Kg.
The difference of CGRP-IR area ratio between the lumbrokinase-treated rats at 600
µg/Kg and the controls reached the significant level at p <0.05.
More macrophages were recruited into the diabetic sciatic fascicles in
lumbrokinase-treated rats (Figure 4). A significantly higher density of macrophages
was noted in the diabetic nerve stumps treated with the lumbrokinase at 600 µg/Kg
compared to the controls and the other two lumbrokinase groups (p < 0.05).
Histological observations showed that myelinated axons were located
dispersedly in the endoneurial areas along with abundant of crescent-shaped Schwann
cells and blood vessels (Figure 5). Mean cross-sectional area of regenerated nerves in
the three groups of lumbrokinase-treated rats was dramatically decreased, showing
less endoneurial edema compared to that in the controls. In addition, myelinated axon
density in the lumbrokinase-treated rats at 600 µg/Kg was significantly larger than
that in the controls (p < 0.05).
Finally, it was found that the IL-1, NGF, PDGF, and TGF-β-mRNA levels were
increased in the diabetic rat sciatic nerve segments after addition of lumbrokinase,
especially both the IL-1 and the PDGF-mRNA reached the significant level at p <
0.05 as compared to the control (Figure 6).
Discussion
Peripheral nerve regeneration involves a complex sequence of events in which axon
regrowth and remyelination of the regenerated axons by Schwann cells are required
for function recovery of the injured nerve (Zhang and Yannas, 2005). Work that has
been done, on diabetic neuropathic nerve, has suggested that down-regulation of some
neurotrophic factors could occur, hindering Schwann cell differentiation, proliferation,
and remyelination (Apfel, 1999). In addition, it has been reported that delay in
macrophage invasion and their later recruitment in diabetic nerves, indicating that
impaired regeneration might be abnormal macrophage participation in nerve repair
(Terada et al., 1998). Also, hyperglycemia-induced blood flow reduction has been
found is an important factor underlying nerve conduction deficits in diabetic
neuropathy (Cameron et al., 1991).
Application of TCM as a means to accelerate the process of tissue and organ
recovery is a new approach (Jiang et al., 2013; Shin et al., 2013; Zhang and Zhao,
2014). Many TCM medications have been used to treat diabetic peripheral neuropathy,
trying to promote nerve repair and regeneration (Ren and Zuo, 2012). However, most
of these medications were extracts from herbs. Recently, more attention has been paid
to the studies relating to the beneficial role of invertebrates’ extracts used in
regenerative medicine. My group has successfully demonstrated that the crude
extraction of earthworm not only could enhance neurite outgrowth from PC12 cells,
but also promote regeneration of myelinated axons in rats (Chen et al., 2010a). We
also found that the extraction of earthworm could stimulate Schwann cell migration
via activation of extracellular matrix-degrading proteolytic enzymes (PAs and
MMP2/9) mediated through mitogen-activated protein kinases ERK1/2 and p38
(Chang et al., 2011). In the present study, we further studied the effects of
lumbrokinase, a group of fibrinolytic enzymes extracted from earthworm (Mihara et
al., 1991), on peripheral nerve regeneration in diabetic rats.
It has been found that myelinated fiber density and number in uninjured nerves
were not influenced by diabetes (Kennedy and Zochodne, 2000). However, it has been
established that the sciatic nerve became stiff in the diabetic rats, resulting in decrease
of blood perfusion in the nerve (Chen et al., 2010b). The other view suggests that
regional ischemia of the main arteries in the limb could result in inhibition of nerve
regenerative processes (Honcharuk et al., 2005). The present study showed that
administration of lumbrokinase in the diabetic rats could induce a transient rise in
their skin perfusion, which was beneficial to restore the nerve regenerative processes.
CGRP is a 37 amino acid neuropeptide that is widely distributed in the central
and peripheral nervous systems, including dorsal root ganglion cells and the neurites
of sensory primary afferents and motorneurons in the spinal cord (Miki et al., 1998).
It has been shown that CGRP-IR was increased in the dorsal root ganglia and laminae
I-V of the spinal dorsal horn after peripheral nerve injury (Zheng et al., 2008). CGRP
is also a vasodilator neuropeptide (Hashikawa-Hobara et al., 2012). Deficits in the
action of CGRP in diabetics may render an ischaemic state in microvessels, which
could hinder the regeneration of injured nerve (Kennedy and Zochodne, 2000). In the
present study, we found that an increased expression of CGRP-IR in dorsal spinal
cords was evident in rat diabetes in response to lumbrokinase, especially at higher
dosages of 600 and 1200 µg/Kg. This finding indicates that the administration of
lumbrokinase has an impact on CGRP expression in diabetic rats and that the response
may exert positive effects on regenerating nerve fibers by providing them a
hyperaemic and trophic growth environment. In addition, CGRP has been recognized
as a neurotrophic peptide that could promote neuromuscular development and
regeneration in the transected hypoglossal nerve (Blesch and Tuszynski, 2001). This
may explain why the present CGRP up-regulation result could lead to a better
electrophysological recovery in the lumbrokinase-treated diabetic rats since the
trophic peptide CGRP could promote more motor axonal growth and improve the
performance of innervated muscle fibers.
The role of macrophages played in peripheral nerve injury is implicated in both
exacerbating and repairing tissue damage at the injury site. Slowed Wallerian
degeneration caused by impairment in the invasion of macrophages has been
considered the major reason of failed regeneration in diabetic nerves (Chen et al.,
2010c). In the present study, a comparatively increased number of macrophages were
observed in lumbrokinase-treated nerves. We also found that lumbrokinase could
stimulate the diabetic nerve segments, releasing more cytokines and neurotrophic
factors, including interleukin-1, nerve growth factor, plate-derived growth factor, and
transformation growth factor-ß, which are beneficial to regenerating nerve fibers. It
has been reported that macrophages could secrete cytokines and neurotrophic factors
during nerve regeneration (Lindholm et al., 1987; Nathan, 1987). Taken together,
these findings imply that the increased macrophages observed in lumbrokinase-treated
rats could produce a speedup in Wallerian degeneration and secrete more nerve
growth-promoting substances following nerve injury, leading to enhancement of the
regenerative response of diabetics. In our future studies, we will try to investigate the
mRNA level of macrophage markers, such as IFN-γ, IL-4, IL-10, and IL-13 to show
the direct correlation between macrophage phenotype and the regeneration outcome in
injured nerves.
In the morphometric comparisons, we found that the density of myelinated axons
in the mid-portion of regenerated sciatic nerves was dramatically increased in the
high-dose lumbrokinase groups compared to that in the control group. The exact
mechanism by which lumbrokinase administration affected the observed results is
uncertain. However, we believe that the nerve growth-promoting effects of
lumbrokinase could be via the pathways known to be associated with aforementioned
results, i.e., accelerated circulatory blood flow, increased expression of CGRP, or
improved macrophage infiltration in the diabetic nerves.
Finally, it was found the nerve growth-promoting effects in the group of
lumbrokinase at 1200 µg/kg were somewhat attenuated as compared to those in the
group of lumbrokinase at 600 µg/kg. Specifically, the regenerated nerves in the group
of lumbrokinase at 1200 µg/kg had less mature nerve morphology, less macrophage
invasion, and poorer electrophysiological performance. This result is similar to that of
Gallo et al. who found the responses of cultured chick dorsal root ganglion neuronal
growth cone to NGF-coated polystyrene bead were prevented as elevating the
background NGF concentration. Boyd and Gordon also showed axonal regeneration
could be inhibited by the administration of high doses of brain-derived neurotrophic
factor by functional blockade of p75 NTR receptors. Similarly, Mohiuddin et al. and
Hirata et al. reported that excessive nerve growth-promoting substances could
suppress the axotomy-induced elevation of growth-associated protein 43 (GAP-43),
resulting in inappropriate reestablishment of injured nerve. Therefore, we believe that
the dosage of lumbrokinase at 1200 µg/kg could be too excessive that may provoke
some adverse responses to the recovery of regenerated nerves.
Conclusions
Results provided the evidence that lumbrokinase could promote regrowth of diabetic
axons possibly via increasing blood circulation, CGRP expression, and macrophage
recruitment. To the best of our knowledge, this is the first study of lumbrokinase’s
effects on peripheral nerve regeneration in diabetic rats. The information from this
study provides a basis to consider using lumbrokinase in clinical trials for diabetic
patients suffering from peripheral nerve injury.
Acknowledgements
Han-Chung Lee and Chun-Hsu Yao contributed equally to this work. This study was
supported by China Medical University under the Aim for Top University Plan of the
Ministry of Education, Taiwan. The authors would also like to thank National Science
Council of the Republic of China, Taiwan (NSC102-2221-E-039-007-MY3), China
Medical University Hospital (DMR-101-097), and Taiwan Ministry of Health and
Welfare
Clinical
Trial
and
Research
Center
of
Excellence
(MOHW103-TDU-B-212-113002) for financially supporting this research.
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Captions
Figure 1: Effects of lumbrokinase on cutaneous blood flow in the hindpaw of diabetic
rats. *Significant differences between conditions, P<0.05.
Figure 2: Between-groups
comparison
of
electrophysiological
functions
in
regenerated nerves supplying gastrocnemius muscle in diabetic rats treated
with different concentrations of lumbrokinase. *Significant differences
between conditions, P<0.05.
Figure 3: CGRP in the dorsal horn of the spinal cord showing immunopositive fibers
stained with diaminobenzidine (arrows) ipsilateral to the injury in diabetic
rats treated with different concentrations of lumbrokinase. *Significant
differences between conditions, P<0.05. Scale bar = 200 µm.
Figure 4: Macrophages stained with CD68 (arrows) in regenerated nerves in diabetic
rats treated with different concentrations of lumbrokinase. *Significant
differences between conditions, P<0.05. Scale bar = 30 µm.
Figure 5: Light imaging of regenerated nerves in diabetic rats treated with different
concentrations
of
lumbrokinase.
*Significant
differences
between
conditions, P<0.05. Scale bar = 30 µm.
Figure 6: Changes in mRNA levels of IL-1, NGF, PDGF, and TGF-β of diabetic rat
sciatic nerve segments conditioned by lumbrokinase.
control.
Table 1: PCR primers
*
P <0.05 versus
Table 1
rat IL1b-F
GCACCTTCTTTTCCTTCATCTTTG
rat IL1b-R
TGCAGCTGTCTAATGGGAACAT
rat NGF-F
GTGGACCCCAAACTGTTTAAGAA
rat NGF-R
AGTCTAAATCCAGAGTGTCCGAAGA
rat TGFb-F
CACCGGAGAGCCCTGGATA
rat TGFb-R
TCCAACCCAGGTCCTTCCTA
rat PDGFa-F
AGGATGCCTTGGAGACAAACC
rat PDGFa-R
TCAATACTTCTCTTCCTGCGAATG
rat-GAPDH-F
GGTGGACCTCATGGCCTACA
Rat-GAPDH-R
CAGCAACTGAGGGCCTCTCT
Day 0
Perfusion unit (pu)
180
160
140
120
100
80
60
40
20
0
A (0)
B (300)
C (600)
D (1200) ug/kg
C (600)
D (1200) ug/kg
Day 14
*
160
*
*
Perfusion unit (pu)
140
120
100
80
60
40
20
0
A (0)
B (300)
Day 28
*
*
160
*
Perfusion unit (pu)
140
*
*
120
100
80
60
40
20
0
A (0)
B (300)
Figure 1
C (600)
D (1200) ug/kg
NCV
*
40.0
*
35.0
30.0
m/s
25.0
20.0
15.0
10.0
5.0
0.0
A (0)
B (300)
C (600)
D (1200) ug/kg
Latency
*
1.6
1.4
1.2
ms
1.0
0.8
0.6
0.4
0.2
0.0
A (0)
B (300)
C (600)
D (1200) ug/kg
Amplitude
*
12.0
10.0
mV
8.0
6.0
4.0
2.0
0.0
A (0)
B (300)
C (600)
D (1200) ug/kg
MAP area
*
*
14.0
12.0
mVms
10.0
8.0
6.0
4.0
2.0
0.0
A (0)
B (300)
Figure 2
C (600)
D (1200) ug/kg
A (0 µg/kg)
B (300 µg/kg)
C (600 µg/kg)
D (1200 µg/kg)
Area ratio
*
3.00%
*
2.50%
2.00%
1.50%
1.00%
0.50%
0.00%
A (0)
B (300)
Figure 3
C (600)
D (1200) ug/kg
A (0 µg/kg)
B (300 µg/kg)
C (600 µg/kg)
D (1200 µg/kg)
Macrophage density
*
*
5000
*
*
#/mm2
4000
3000
2000
1000
0
A (0)
B (300)
Figure 4
C (600)
D (1200) ug/kg
A (0 µg/kg)
B (300 µg/kg)
C (600 µg/kg)
D (1200 µg/kg)
Cross-sectional area
0.25
mm2
0.2
0.15
0.1
0.05
0
A (0)
B (300)
C (600)
D (1200) ug/kg
Axon density
*
25000
*
*
*
#/mm2
20000
15000
10000
5000
0
A (0)
B (300)
Figure 5
C (600)
D (1200) ug/kg
Gene Expression
*
RQ (Relative Quantification)
1.8
1.6
*
1.4
1.2
1
0.8
0.6
0.4
0.2
0
CONTORL
IL1b
NGF
Figure 6
PDGFa
TGFb