J Neuropathol Exp Neurol
Copyright Ó 2011 by the American Association of Neuropathologists, Inc.
Vol. 70, No. 5
May 2011
pp. 323Y339
ORIGINAL ARTICLE
Motor End Plate Innervation Loss in Diabetes
and the Role of Insulin
George J. Francis, BSc, Jose A. Martinez, MSc, Wei Q. Liu, MSc, Douglas W. Zochodne, MD,
Leah R. Hanson, PhD, William H. Frey, II, PhD, and Cory Toth, MD
Abstract
Retraction of distal sensory axons is a prominent feature in diabetic
peripheral neuropathy (DPN), a process amenable to insulin therapy.
Nevertheless, diabetic patients and long-term diabetic mice develop
motor deficits after longer durations of DPN, a process that may be
related to insulin deficiency. To compare the efficacy of intranasal
delivery of insulin (IN-I) and subcutaneous insulin (Subc-I) in preventing motor deficits in a long-term mouse model of DPN, IN-I or
Subc-I, 0.87 IU daily or placebo was delivered in separate cohorts of
diabetic and nondiabetic CD1 mice for 8 months. Radiolabeled detection was used to assess insulin delivery and biodistribution. Biweekly
behavioral tests and monthly electrophysiological and multipoint
quantitative studies assessed motor function deficits. Morphometric
analysis of spinal cord, peripheral nerve, muscle innervation, and specific molecular markers were evaluated at and before the end point.
Despite progressive distal axonal terminal loss, numbers and caliber
of motor neurons were preserved. There were no differences in glycemia between IN-I and Subc-I-treated mice. Intranasal delivery of
insulin and, to a lesser extent, Subc-I, protected against electrophysiological decline, loss of neuromuscular junctions, and loss of motor
behavioral skills. Intranasal delivery of insulin was associated with
greater preservation of the phosphatidylinositol 3-kinase signaling
pathway involving Akt, cyclic AMP response element binding protein, and glycogen synthase kinase 3A but did not alter extracellular
signalYregulated kinase, mitogen-activated protein kinase/extracellular
signalYregulated kinase, or c-Jun amino-terminal kinase. Thus, direct
delivery of insulin to the nervous system might prevent motor deficit
in human type 1 diabetes by preservation of the phosphatidylinositol
3-kinaseYAkt pathway rather than only affecting glycemic levels; the
effects of insulin on other signaling pathways may, however, play
additional roles.
Key Words: Akt, Denervation, Diabetes, Insulin, Motor neuron,
Neuropathy, PI3K.
INTRODUCTION
From the Department of Clinical Neurosciences and the Hotchkiss Brain
Institute (GJF, JAM, WQL, DWZ, CT), University of Calgary, Calgary,
Alberta, Canada; and Alzheimer’s Research Center at Regions Hospital
(LRH, WHF), HealthPartners Research Foundation, St. Paul, Minnesota.
Send correspondence to: Cory Toth, MD, Department of Clinical Neurosciences, University of Calgary, Room 155, 3330 Hospital Dr, NW,
Calgary, Alberta, Canada T2N 4N1; E-mail: [email protected]
This work was funded by the Alberta Heritage Foundation for Medical
Research (AHFMR). Dr. C. Toth is a clinical investigator of AHFMR.
Dr. D. Zochodne is a scientist of AHFMR. Dr. W. Frey II has an intellectual property patent on the use of intranasal insulin administration for
neurologic agents (Frey WH 2nd. Neurologic agents for nasal administration to the brain. World Intellectual Property Organization. PCT
priority date 5.12.89, WO 91/07947. June 13, 1991). The investigators have
no financial interest in monetary profit with performance of these studies.
Supplemental digital content is available for this article. Direct URL citations
appear in the printed text and are provided in the HTML and PDF versions
of this article on the journal’s Web site (www.jneuropath.com).
Diabetes mellitus in humans commonly leads to development of diabetic peripheral neuropathy (DPN), a diffuse
disorder of peripheral nerves with ‘‘stocking and glove’’ loss
of sensation despite initially preserved distal motor function
(1). In 70% of patients with DPN, there is a mixed pattern of
sensory, autonomic, and motor disturbance (1); these may
lead to weakness and muscle atrophy of distal leg and foot
muscles (2Y4) and contribute to falling (5).
Information regarding motor neuron populations in diabetic models has been modest; short-duration models have
not identified problems with myocontractility or other motor
deficits beyond conduction slowing (6, 7). However, later
stages of diabetes may be associated with a form of motor
neuropathy or motor neuronopathy. Mice with experimental
streptozotocin (STZ)-induced DPN develop motor nerve conduction velocity (MNCV) slowing, loss of compound motor
action potential (CMAP) amplitude, and an electrophysiological loss of motor units (7, 8). Increased single motor unit
action potential (SMUP) in diabetes (8) suggests either an
enlargement of motor units through compensatory sprouting or
preferential loss of smaller motor units (9). An estimated loss
of motor units has been demonstrated at relatively early stages
of STZ-induced diabetes in mice; this is associated with a
loss of ionic current amplitudes and fragmentation of clusters
of acetylcholine receptors at the motor end plate (10). These
changes suggest early remodeling of motor units during DPN.
Such changes are overshadowed in human patients in which
sensory manifestations dominate, but motor deterioration does
occur in later stages of human DPN (4, 11, 12).
Current theories regarding the development of DPN
include effects of chronic hyperglycemia (13, 14), excessive
sorbitol-aldose reductase pathway flux (15), overactivity of
protein kinase C isoform(s) (16), increased oxidative and
nitrergic stress (17), microangiopathy (18), neurotrophin deficiency (19, 20), and a role for advanced glycation end products
and their receptor (7, 21). In addition, the impaired availability,
action, or uptake of insulin and insulin-like growth factor-1,
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
323
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Francis et al
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
both of which are neurotrophic factors, may contribute to
neuronal dysfunction (22), particularly within sensory neurons
(23Y25). Insulin receptor (IR) binding promotes tyrosine autophosphorylation of the IRA subunit, leading to subsequent
phosphorylation of cellular substrates, including IR substrate-1
(IRS-1). This sequence creates an active signaling complex that
involves phosphatidylinositol 3 kinase (PI3K), Akt, and the
downstream effectors cyclic AMP response element binding
protein (CREB) and glycogen synthase kinase 3A (GSK-3A),
among other molecules (26, 27). The presence of IRs at ventral
horn cells suggests that an insulin neurotrophic deficit may
contribute to the development of diabetic motor neuropathy.
We hypothesized that intranasal insulin delivery (IN-I)
would slow development of motor impairment through downstream effects on the distal-most end plate terminal muscle
innervation. Intranasal delivery targets insulin to the nervous
system, thereby bypassing the blood-brain barrier and without
significantly altering blood levels of insulin or glucose (28).
This paradigm permits separate evaluation of the trophic
versus the antihyperglycemic effects of insulin on the nervous
system. Here, we compared the effects of IN and similar
amounts of Subc insulin in an experimental type 1 diabetes
model of DPN.
assessment. Mice with diabetes are indicated with a ‘‘D’’ and
mice without diabetes (control mice) are indicated with a ‘‘C.’’
Delivery of subcutaneous saline is indicated as ‘‘Subc-S’’;
subcutaneous insulin is indicated as ‘‘Subc-I’’ (Fig. 1).
Pharmacokinetic studies of IN or Subc delivery of
125
I-labeled insulin have been performed previously in mice
at the University of Minnesota to define insulin concentrations at tissues of interest (29, 30). Experiments described
herein were performed in mice concurrently studied for sensory nerve dysfunction (30).
Daily IN-I and Subc-I Delivery
Daily IN-I (Humulin R; Eli Lilly, Toronto, Ontario,
Canada) and IN-S were administered to either diabetic or
nondiabetic male CD1 mice after a 1-week training period
using only IN-S for accustoming the mice before diabetes
verification. While held in supine position and in neck extension, a total of 24 KL containing either 0.87 IU of insulin
or 0.9% saline only was given to each mouse as 4 drops of
6 KL each provided by Eppendorf pipette over alternating
nares every 1 minute. Daily Subc-I (Humulin R) or Subc-S
was also administered daily to mice at the same dose. Subc
injections of either insulin or saline occurred over the lower
MATERIALS AND METHODS
Animals
A total of 491 male CD1 wild-type mice with initial
weights of 20 to 30 g were housed in plastic sawdust-covered,
pathogen-free cages with a normal light-dark cycle and free
access to mouse chow and water. All protocols were reviewed
and approved by the University of Calgary Animal Care
Committee using the Canadian Council of Animal Care
guidelines; principles of laboratory animal care were strictly
followed. At 1 month, 332 mice were injected with STZ
(Sigma, St. Louis, MO) intraperitoneally once daily for each of
3 consecutive days with doses of 60, 50, and then 40 mg/kg.
The remaining 159 mice were injected with placebo (sodium
citrate) for 3 consecutive days. A total of 16 mice were harvested to obtain measurements of insulin in the hours after
IN-I or intranasal saline (IN-S). A total of 156 mice injected
with STZ and 80 mice injected with carrier or IN-I (6 mice)
were designated for studies performed after 1, 3, and 5 months
of diabetes; the remaining mice were followed for the entire
length of the study (i.e. 8 months of diabetes or equivalent for
carrier-injected mice), as mortality permitted. A selection of 12
mice received IN-I only after 8 months of untreated diabetes for
a total of 2 days (acute therapy) or 7 days (subacute therapy).
Monthly whole-blood glucose measurements were performed
using the tail vein and a blood glucometer (OneTouch Ultra
Meter; LifeScan Canada, Burnaby, British Columbia, Canada).
Hyperglycemia was verified 1 week after STZ injections with
tail vein sampling of fasting whole blood, with a glucose level
of 16 mmol/L or higher (reference range, 5Y8 mmol/L) required
for diagnosis of diabetes. All animals had whole-blood glucose sampling and weight calculations monthly. Mice were
followed and harvested at 1, 3, 5, or 8 months of diabetes (up
to the age of 9 months), except for mice that did not develop
diabetes as defined above; these were excluded from further
324
FIGURE 1. Flowchart of all diabetic and nondiabetic mice
throughout the described studies. Mice received either streptozotocin (STZ) or carrier (placebo) at age 1 month, followed
by verification of diabetes 1 week later. After an additional
week, at 1.5 months of age, intranasal (IN) or subcutaneous
(Subc) delivery of insulin (I) or saline (S) was initiated. Twicemonthly behavioral testing and once-monthly blood glucose
and electrophysiological testing were performed. Harvesting
end points occurred after 1, 3, 5, or 8 months of diabetes.
Numbers under the corresponding arrow lines represent the
numbers of mice in each cohort harvested at each end point of
2, 4, 6, and 9 months of age. At age 9 months, the numbers of
survivors (numerator) and numbers initially in each long-term
cohort (denominator) are shown.
Ó 2011 American Association of Neuropathologists, Inc.
Copyright © 2011 by the American Association of Neuropathologists, Inc. Unauthorized reproduction of this article is prohibited.
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Motor Unit Loss in Diabetes
torso using rotating locations. All treatments began immediately after confirmation of the presence of diabetes for each
cohort at 1 week after injections of STZ/citrate concluded. In
the first week, daily glucometer testing was performed for
all mice, followed by once-monthly testing. On the day of
harvesting, the last saline or insulin dose was administered
6 hours before death.
was placed on the pull rod, with digits encircling the rod. For
combined forelimb testing, both forelimbs were placed on the
pull rod with all digits of each forepaw allowed to encircle the
rod. Peak strength (g) was recorded for each hind limb and for
the combined forelimbs, with the top 3 results of 5 consecutive
trials recorded. Rearing was performed with individual mice
placed in a closed arena. Scoring was given each time a mouse
reared (raised its forepaws off the floor) with or without wall
support, in either the periphery or in the center of the test box,
within a 10-minute interval. For all motor testing, the investigator was blinded to the treatment received by each mouse.
Electrophysiology During IN-I and Subc-I
Delivery Studies
Electrophysiological assessment of sciatic nerve conduction was performed as previously described (7, 31) under
halothane anesthesia. Initial baseline studies before STZ or
placebo injections revealed no significant differences between
cohort groups. For all groups, 6 to 8 mice underwent electrophysiological testing before induction of diabetes and after
each month of diabetes. All stimulating and recording electrodes were platinum subdermal needle electrodes (Grass
Instruments, Astro-Med, West Warwick, RI), with near-nerve
temperature kept constant at 37 T 0.5-C using a heating lamp.
For orthodromic motor studies, an active recording electrode was placed in the peroneal nerveYinnervated extensor
digitorum brevis muscle, with a reference electrode placed
at the dorsal aspect of the first metatarsophalangeal joint.
Stimulation electrodes were placed at either the sciatic notch
or the popliteal fossa for sciatic or tibial nerve stimulations,
respectively. Both the CMAP amplitude and MNCV were
measured. Single motor unit action potential amplitudes
were also measured monthly by eliciting the first all-or-none
potentials over threshold with the smallest injection of current. Motor unit number estimation (MUNE) was performed
using an incremental method (31). In brief, stimulus intensity
was slowly increased from subthreshold levels until a small,
all-or-none response was evoked (initial SMUP amplitude),
with consistent amplitude and appearance on 3 separate occasions without any smaller amplitude responses evoked.
Stimulation intensity was then slowly increased providing
identification of stepwise quantal increments in amplitude.
This was repeated for a minimum of 25 increments, unless
fewer incremental steps were not obtainable. Individual motor
unit amplitudes were estimated by subtraction of amplitudes
of each response from the prior response. An average of individual values was used to estimate the average SMUP
amplitude; individual values differing by 2 SDs greater than
or less than the average SMUP were discarded. The resulting
average SMUP was divided into the supramaximal CMAP
amplitude response, yielding the MUNE.
Behavioral Testing During IN-I and Subc-I
Delivery Studies
Behavioral testing to evaluate hind limb/forelimb strength
was performed twice monthly on 10 mice in each cohort. A
2-week training period was performed to accustom mice to
each procedure before diabetes verification. A Chatillon DFIS-2
digital force measurement dynamometer (Ametek, Inc., Paoli,
PA) was used to perform quantification of individual hind limb
and combined forelimb strength testing. Peak values were
measured using a sampling rate of 1 kHz. Mice were held by the
posterior cervical region with hind limbs free as 1 hind limb
Harvesting of Nervous Tissues After IN-I and
Subc-I Delivery Studies
After 1, 3, 5, or 8 months of diabetes, mice from each
cohort were killed (Fig. 1). Euthanasia and harvesting of
tissues were performed 6 hours after the last IN or Subc
delivery of insulin or saline under pentobarbital (60 mg/kg)
anesthesia. A 0.5-mL volume of whole intracardiac blood
was used for glycated hemoglobin A1c measurements performed with affinity chromatography (Calgary Laboratory
Services, Calgary, Alberta, Canada). The following tissues
were harvested: cervical, thoracic and lumbar spinal cord,
and bilateral sciatic nerves, proximal and distal portions of
peroneal nerves, and tibialis anterior and extensor digitorum
muscles. One half of all tissues (left side) were placed either
in Zamboni fixative for later immunohistochemistry or were
fixed in cacodylate-buffered glutaraldehyde, then cacodylate
buffer for later Epon embedding (Epon 812 resin; Canemco,
Inc., Lakefield, PQ, Canada) for morphometric studies. The
remaining (right side) tissues were immediately fresh frozen
at j80-C or placed in TRIzol (Invitrogen, Burlington,
Ontario, Canada) and stored at j80-C for protein and RNA
investigations, respectively. Small portions of fresh frozen
cerebrum and cervical spinal cord were used to determine
insulin concentrations after 8 months.
For immunohistochemistry, specimens were prepared
using 10-Km cryostat transverse and longitudinal nerve sections, as previously described (30). Application of primary and
secondary antibodies was also performed as described (30).
Tissue Insulin Concentrations, PI3K/Akt Pathway
Protein Phosphorylation Ratios, and Plasma
Glucose and Glycated Hemoglobin A1c
Measurements
For a total of 51 mice, cerebral and cervical spinal cord
samples were obtained to measure insulin concentration using
microparticle enzyme immunoassays (MEIA Insulin, IMX
System; Abbott Laboratories, Chicago, IL) with minimal detectable amount of 0.1 mU/mL. Three nondiabetic mice were
killed after 1 month of age, the time point when STZ was to be
delivered, at 1 hour after IN-S delivery. Three C IN-S, 3 D IN-S,
3 D Subc-I, and 3 D IN-I mice were killed at each time point of
2, 5, 7, and 9 months, each at 1 hour after insulin or saline
treatment. Cerebral samples were obtained from the frontal
cortex; cervical spinal cord samples were taken from the entire
cervicomedullary junction. Tissues were placed in 0.5 mL of
phosphate-buffered saline (PBS) and were homogenized at
low speed to form the supernatant.
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Copyright © 2011 by the American Association of Neuropathologists, Inc. Unauthorized reproduction of this article is prohibited.
325
Francis et al
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Blood glucose from tail veins of 3 diabetic and 3 nondiabetic mice after 2 months of age (1 month of diabetes) at 1,
2, 4, 8, and 24 hours after IN and Subc delivery of insulin or
saline was examined. In these mice, lumbar spinal cords at the
same time points in D IN-I, D Subc-I, D IN-S, C IN-I, C
Subc-I, and C IN-S mice were harvested to measure proteins
in the PI3K/Akt pathway.
(Miles Laboratories, Elkhart, IN) and frozen with cold isopentane. Optimum cutting temperature blocks were cut into
3-mm3 cubes, and tissue was then serially sectioned longitudinally at 25 Km and placed on poly-d-lysineYcoated
slides, which were first rinsed with PBS and blocked with
1% goat serum solution. The slides were then incubated
for 24 hours at 4-C with rabbit antiserum to neurofilament
(NF-200, 1:200; Chemicon International, Temecula, CA).
Slides were then washed with PBS and incubated with
fluorescent secondary antibody goatYanti-rabbitYAlexa 488
(1:400; Chemicon) and >-bungarotoxinYtetramethyl rhodamine conjugate (1:500; Molecular Probes, Invitrogen) for
1 hour at room temperature. After further washing in PBS,
slides were mounted with bicarbonate-buffered glycerol,
coverslipped, and viewed under a fluorescence microscope
(Zeiss Axioskope; Zeiss) and confocal microscope (Olympus
FV 300 Confocal Microscope; Olympus, Melville, NY).
Images were analyzed at 200 magnification, with
selected images obtained using confocal laser microscopy
(Nikon, Inc., Melville, NY). Motor end plates were visualized
in red by labeling with Cy3-conjugated >-bungarotoxin. It
was then determined whether the motor end plate was a free
end plate (no visible axon attachment, classified as ‘‘0’’) or an
innervated end plate. Innervated end plates were scored using
the following scheme: (1a) ultraterminal sprout, (1b) preterminal sprout, (1c) nodal sprout, and (2) terminal nerve
ending (i.e. not innervated by a sprouted axon) (9). Ultraterminal sprouts grow from intact axons at the motor end
plates, preterminal sprouts arise from nerve terminals, nodal
sprouts arise from nodes of Ranvier, and ultraterminal and
preterminal sprouts are generally both referred to as terminal
sprouts (9). The total number of motor end plate profiles and
the innervation scores from each muscle were calculated, and
the frequency of each innervation score was compared
between cohort groups, as previously described (9).
We also took fresh frozen sections from extensor digitorum brevis muscles that were cut at 10 Km using a cryostat
from selected D IN-S and C IN-S mice. Sections were examined using standard protocols for ATPase histochemical
staining to distinguish fiber types using a pH solution of 10.2
(33). These sections were visually inspected for evidence of
myopathic or neuropathic changes.
Quantitative Morphometry of Peripheral Nerve
and Spinal Cord From Mice Administered IN-I
and Subc-I
For peripheral nerve and spinal cord specimens, Eponembedded samples were cut by an ultramicrotome at 1 Km
and stained with 0.5% toluidine blue (7). Additional spinal
cord specimens were stained with Cresyl violet. Image analysis was performed by a single examiner blinded to the origin
of the sections (Zeiss Axioskope [Zeiss, Toronto, Canada] at
400 and 1,000 magnifications using Scion Image v.4.0.2
[Scion, Inc., Fredrick, MD]) with measurements of the number, axonal area, fascicle area, and myelin thickness of all
myelinated fibers within 25 nonadjacent transverse nerve
sections in the sciatic nerve and proximal and distal portions
of peroneal nerve. G ratios were calculated using the following formula: axon area / (axon + myelin area). Axonal fiber
densities were expressed as the number of axons within 1
transverse nerve section divided by the transverse area of the
sampled nerve section. For spinal cord, neurons with visible
nuclei were used for counting within an area sized for 25
nonadjacent sections separated by approximately 300 Km for
each cervical, thoracic, and lumbar spinal cord sample to
measure the neuronal density within the ventral horn and to
provide calculations for the estimated neuronal densities (32).
All measurements of both nerve and cord morphometry were
performed using Scion Image v.4.0.2 (Scion, Inc). For each
mouse, motor neurons in lamina IX were counted at the level
of the cervical (C5Y7), thoracic (T6Y8), or lumbar spinal cord
(L4Y6), with different regions having nonoverlapping neurons
examined. The identification of the cervical, thoracic, and
lumbar tract of the spinal cord was based on the presence of
the corresponding enlargements of the spinal cord diameter at
cervical and lumbar regions. Morphometry assessment consisted of outlining motor neurons from lamina IX in spinal
levels at L4Y6 with a clearly visible neuronal perimeter, excluding the cellular processes, to determine the neuronal area
using Adobe Photoshop 9.0 (Adobe, San Jose, CA). We also
assessed for the degree of vacuolation considered as the
prevalence among individual motor neurons within each tissue section.
Quantitative Morphometry of Neuromuscular
Junctions and Muscle Fiber Typing
To examine the neuromuscular junction (NMJ), the
tibialis anterior and extensor digitorum brevis muscles (on
the side opposite of the electrophysiologically tested side)
were harvested and incubated in Zamboni fixative overnight
at 4-C. The tissues were washed 3 times with 1 mol/L of PBS
5 minutes each and incubated in 20% sucrose solution in
1 mol/L of PBS overnight at 4-C. Tibialis anterior samples
were embedded in optimum cutting temperature compound
326
Immunohistochemistry
Immunohistochemistry was performed as described
previously (27) using primary antibodies to NeuN (1:100;
Chemicon). The PI3K and Akt pathway were investigated
with PI3K (1:200; Santa Cruz Biotechnology, Inc., Santa
Cruz, CA), PKB/Akt (1:200, antiYprotein kinase B [Akt];
Stressgen, Victoria, Canada), pAkt (1:200, antiYphospho-Akt
[1:200], anti-Ser473; Cell Signaling Technologies, Danvers,
MA), and the nuclear signaling transcription factor NF-JB
p65 subunit (1:200, antiYNF-JB p65; Santa Cruz Biotechnology) and p50 subunit (1:200, antiYNF-JB p50; Santa Cruz
Biotechnology). Additional immunohistochemistry was performed for CREB (1:200; Abcam, Inc., Cambridge, MA) and
GSK-3A (1:200; Abcam).
Tissue specimens were examined under fluorescence
microscopy (Zeiss Axioskope, Axiovision, and Axiocam; Zeiss
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Copyright © 2011 by the American Association of Neuropathologists, Inc. Unauthorized reproduction of this article is prohibited.
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Motor Unit Loss in Diabetes
TABLE 1. Insulin Concentrations Within the Cerebrum and Spinal Cord After Intranasal or Subcutaneous Delivery Insulin or
Intranasal Saline Delivery After 1 to 8 Months of Diabetes
Age
Tissue Location
Cerebrum
Cervical spinal cord
Cohort
1 mo
2 mo
5 mo
7 mo
9 mo
C IN-S
D IN-I
D Subc-I
D IN-S
C IN-S
D IN-I
D Subc-I
D IN-S
0.09 T 0.02
0.08 T 0.01*
0.42 T 0.05†
0.11 T 0.02‡
0.01 T 0.00
0.08 T 0.02*
0.12 T 0.02†
0.06 T 0.02‡
0
0.06 T 0.01*
0.39 T 0.04†
0.09 T 0.01‡
0
0.07 T 0.01*
0.13 T 0.02†
0.03 T 0.01‡
0
0.07 T 0.01*
0.36 T 0.05†
0.10 T 0.01‡
0
0.08 T 0.01*
0.08 T 0.01†
0.03 T 0.01‡
0
0.06 T 0.01*
0.40 T 0.03†
0.08 T 0.01‡
0
0.06 T 0.01*
0.10 T 0.02†
0.04 T 0.01‡
0
0.06 T 0.01
All measurements are mean T SEM, with units expressed as milliunits per gram of tissue. All insulin concentrations were determined using microparticle enzyme
immunoassay (MEIA).
*indicates significance with comparison of nondiabetic (control, C) IN-S mice to D IN-S mice; †indicates significance with comparison of diabetic (D) IN-I mice to both D Subc-I
and D IN-S mice; ‡indicates significance with comparison of D Subc-I mice to D IN-S mice using multiple ANOVA testing with Bonferroni post hoc t test comparisons (> = 0.05,
p G 0.016) (nonmatched ANOVA tests, F values range between 1.66 and 12.77 for indicated groups and time points, df = 2, n = 3).
IN-I, intranasal insulin; IN-S, intranasal saline; Subc-I, subcutaneous insulin; Subc-S, delivery of subcutaneous saline.
Stressgen) and pAkt (1:1000, anti-Ser473; Cell Signaling Technologies). The nuclear signaling transcription factor
NF-JB p65 and p50 subunits (1:1000 each) were also examined, and additional Western blotting was performed for
CREB (1:500), phospho-CREB (anti-Ser133, 1:500), GSK3A (1:500), and phosphoYGSK-3A (anti-Ser9, 1:500) (all from
Abcam, Inc.). For housekeeping, antiYA-actin (1:100; Biogenesis Ltd., Poole, UK) was applied to separate blots. For the
assessment of additional important insulin signaling pathways, supplementary blots were probed with MEK (1:1000),
pMEK (1:2000), extracellular signalYregulated kinase (ERK;
1:2000), pERK (1:2000), c-Jun amino-terminal kinase (JNK;
1:1000), and pJNK (1:1000) antibodies (all from Cell Signaling Technologies) for STZ-injected mice that received
2 days, 1 week, or 8 months IN-I. Secondary anti-rabbit, antimouse, or anti-human IgG horseradish peroxidaseYlinked
antibody (Cell Signaling Technologies) was applied at 1:5000
in each case as appropriate. Signal detection was performed
by exposing of the blot to enhanced chemiluminescent
reagents ECL (Amersham, Baie d’Urfe, Quebec, Canada) for
Canada) at 400, and images were obtained based on anatomic location. For spinal cord, each of NeuN, S100, PI3K,
Akt, pAkt, and NF-JB immunolabeled profiles was identified
in conjunction with its cellular profile using Adobe Photoshop.
Anterior horn cell neurons from the cervical, thoracic, or lumbar region were identified within 10-Km-thick sections obtained
through the midportion of the individual spinal cord level only
for visualization purposes.
Western Immunoblotting
Tissue portions were homogenized using a RotorStator
Homogenizer in ice-cold lysis buffer (10% glycerol, 2% SDS,
25 mmol/L Tris-HCl, pH 7.4; Roche Mini-Complete Protease
Inhibitors, Laval, PQ, Canada). Samples were then centrifuged at 10,000 g for 15 minutes, and equal amounts (15 Kg)
of protein were separated by SDS-PAGE using 10% polyacrylamide gels as previously described (7).
Blots were probed with antibodies to IRA (1:500),
IRS-1 (1:500), and PI3K (1:1000) (all from Santa Cruz
Biotechnology) and with antibodies to PKB/Akt (1:1000;
TABLE 2. Venous Blood Glucose Levels in the 24 Hours After Subcutaneous or Intranasal Intervention in Diabetic and
Nondiabetic Mice at 2 Months of Age (1 Month of Diabetes)
Hours After Intervention
Cohort
C IN-I
C IN-S
C Subc-I
C Subc-S
D IN-I
D IN-S
D Subc-I
D Subc-S
j1
1
2
4
8
24
5.9 T 0.6
6.0 T 0.8
5.5 T 0.7
5.2 T 0.5
33.1 T 1.6
33.0 T 1.4
33.2 T 1.1
33.1 T 1.8
6.1 T 0.7
6.4 T 0.9
5.1 T 1.0
6.0 T 0.8
33.0 T 1.5
33.1 T 2.0
24.6 T 1.9†
33.1 T 2.3
5.5 T 0.6
6.0 T 0.7
3.2 T 1.1*
5.3 T 0.7
32.8 T 1.3
33.0 T 1.6
22.8 T 2.3†
33.1 T 2.1
5.4 T 0.8
5.6 T 1.0
2.8 T 1.1*
5.4 T 0.7
32.9 T 1.5
33.1 T 1.9
29.3 T 1.7
33.0 T 2.1
5.3 T 0.8
6.1 T 0.5
3.9 T 0.9
6.0 T 0.7
33.0 T 1.8
33.0 T 2.1
32.1 T 2.2
32.8 T 2.0
6.0 T 0.7
6.0 T 0.8
4.8 T 1.1
4.8 T 0.9
33.1 T 1.8
33.2 T 1.1
32.9 T 2.0
33.1 T 2.1
All measurements are mean T SEM, with units expressed as millimoles per liter.
Multiple nonparametric Kruskal-Wallis ANOVA tests were performed with Bonferroni post hoc t test comparisons (> = 0.05, p G 0.016).
*indicates significance with comparison of non-diabetic (control, C) IN-I mice to C IN-S mice; †indicates significance with comparison of diabetic (D) IN-I mice to D Subc-I mice;
(nonmatched ANOVA tests, F values range between 1.12 and 2.51 for indicated groups and time points, df = 2, n = 3).
C, control mice; D, diabetic mice; IN-I, intranasal insulin; IN-S, intranasal saline; Subc-I, subcutaneous insulin; Subc-S, delivery of subcutaneous saline.
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Francis et al
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
2 minutes. The blots were subsequently exposed and captured on Kodak X-OMAT K film (Sigma Inc., St. Louis,
MO). In each case, 3 blots were performed from each cohort
of diabetic and nondiabetic mice, with analysis performed
using Adobe Photoshop 9.0 for quantification of blotting
densities.
Primer sequences for PI3K were the following: forward, 5¶AACCCGGCACTGTGCATAAA-3¶; and reverse, 5¶-GCCCA
TTGGATTAGCATTGATG-3¶. Primer sequences for Akt
were the following: forward, 5¶-TCTGCCCTGGACTACTTG
CACT-3¶; and reverse, 5¶-GCCCGAAGTCCGTTATCTTGA3¶. Primer sequences for NF-JB p65 were the following: forward, 5¶-TGTGCGACAAGGTGCAGAAA-3¶; and reverse,
5¶- ACAATGGCCACTTGCCGAT-3¶. 18S primers and probe
sequences were the following: forward, 5¶-TCCCTAGTGAT
CCCCGAGAAGT-3¶; and reverse, 5¶-CCCTTAATGGCAGT
GATAGCGA-3¶. Reverse transcriptionYpolymerase chain reaction was done using SYBR Green dye. All reactions were
performed in triplicate in an ABI PRISM 7000 Sequence
Detection System (Applied Biosystems Canada, Streetsville,
ON, Canada). Data were calculated by the 2j$$CT method and
presented as the fold induction of messenger RNA (mRNA) for
the gene of interest in diabetic tissues normalized to the
housekeeping gene 18S compared with nondiabetic tissues
(defined as 1.0-fold).
Messenger RNA Quantification
Total RNA was extracted from peripheral nerve and
spinal cord using TRIzol (Invitrogen). Total RNA (1 Kg) was
processed directly to complementary DNA synthesis using
the SuperScript II Reverse Transcriptase system (Invitrogen).
Analysis
All data are presented as mean T SEM. For blood glucose comparisons, where values greater 33.3 mmol/L could
not be detected, nonparametric testing using Kruskal-Wallis
1-way analysis of variance (ANOVA) testing was performed.
Analysis of variance testing with multiple comparisons of
independently assessed samples and groups was performed
in all other cases. For serially collected electrophysiological
and behavioral data, repeated-measures ANOVA testing was
performed. Bonferroni corrections were applied in all cases
where multiple groups were examined. All statistical comparisons were intended between the following groups only: D
IN-I and D Subc-I, D IN-I and D IN-S, D IN-I and C IN-I, D
Subc-I and D Subc-S, D Subc-I and C Subc-I, C IN-I and C
Subc-I, C IN-I and C IN-S, and C Subc-I and C Subc-S. For
studies of short-term IN or Subc intervention, neighboring
time points were compared with ANOVA.
FIGURE 2. Motor behavioral data for mice with [D] or without
[C] diabetes. (A) Loss of rearing ability was first seen in diabetic
mice after 5 to 7 weeks of diabetes. Delivery of subcutaneous
saline is indicated as ‘‘Subc-S,’’ subcutaneous insulin as ‘‘Subc-I,’’
intranasal saline as ‘‘IN-S,’’ and intranasal insulin as ‘‘IN-I.’’ Both
D IN-I and D Subc-I mice had significant amelioration of lost
rearing ability after 11 weeks of testing, but only D IN-I mice
maintained this benefit consistently over the remainder of the
testing period. (B, C) Both hind limb (B) and combined forelimb grip strength testing (C) showed deficit in diabetic mouse
cohorts after 5 to 7 weeks, whereas D IN-I mice showed protection against loss of grip strength after 19 to 21 weeks of
diabetes for both hind limb (B) and combined forelimb (C)
grip strength that was maintained until the end of the testing.
Significant differences were determined by multiple repeatedmeasures ANOVA tests, with *indicating significant difference
(> G 0.05, p G 0.016 using Bonferroni corrections) between the
D IN-I mouse group and other diabetic mouse cohorts, and
?
indicating significant difference (> G 0.05, p G 0.016 using
Bonferroni corrections) between the D Subc-I mouse group
and D Subc-S and D IN-S groups for the respective time points
(n = 6Y8 mice in each mouse cohort for each time point).
328
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J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
RESULTS
Diabetes Model
After STZ injection, 262 (86%) of 304 mice developed
diabetes within 2 weeks, which was maintained during the
length of the study. Diabetic mice were smaller than nondiabetic
mice at 5 and 8 months after STZ injection and also had smaller
body weights 5 or 8 months of diabetes (Table, Supplemental
Digital Content 1, http://links.lww.com/NEN/A224), as previously described (30). D IN-I mice maintained weight better
Motor Unit Loss in Diabetes
than the cohort IN-S mice. Hyperglycemia was identical in
mice receiving IN-I or IN-S, but Subc-I mice had more documented hypoglycemia and more episodes of illness or death
associated with hypoglycemia. Glycated hemoglobin A1c was
increased in all diabetic mice at 8 months of diabetes and
was identical between D IN-I and D IN-S mice; it was reduced
in surviving D Subc-I mice. The mortality rate in diabetic
mice was significantly higher than in nondiabetic mice, although D IN-I mice had less mortality relative to D IN-S, D
Subc-S, and D Subc-I mice (Table, Supplemental Digital
FIGURE 3. Electrophysiological testing of sciatic nerves in mice with [D] or without [C] diabetes demonstrated motor impairment.
Delivery of subcutaneous saline is indicated as ‘‘Subc-S,’’ subcutaneous insulin as ‘‘Subc-I,’’ intranasal saline as ‘‘IN-S,’’ and intranasal insulin as ‘‘IN-I.’’ (A, B) Beginning at approximately 3 months of diabetes, D IN-I mice had amelioration of motor decline of
compound motor action potential (CMAP) amplitudes (A) and motor nerve conduction velocities (MNCV) (B) after 5 months of
diabetes versus other diabetic cohorts. Diabetic Subc-I mice had less significant prevention of diabetes-mediated decline. (C, D)
Protection against motor unit loss was only identified for D IN-I mice when compared with other diabetic mice during motor unit
number estimation (MUNE) testing after 6 months of diabetes. (C) Diabetic mice demonstrated motor unit loss after 2 to 3 months
of diabetes. The initial single motor unit action potential (SMUP) amplitude size demonstrated age-dependent increases in nondiabetic mice (D), with magnified increases noted in diabetic mice. Both D IN-I and D Subc-I mice demonstrated protection against
increasing initial SMUP amplitude size after 5 to 6 months of diabetes, with more definitive protection in D IN-I mice. Significant
differences were determined by multiple repeated-measures ANOVA tests; *indicates significant difference (> G 0.05, p G 0.016
using Bonferroni corrections) between the D IN-I mouse group and other diabetic mouse cohorts; ?, significant difference (> G 0.05,
p G 0.016 using Bonferroni corrections) between the D Subc-I mouse group and diabetic Subc-S and diabetic IN-S groups for the
respective time points (n = 6Y8 mice in each mouse cohort for each time point).
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329
Francis et al
Content 1, http://links.lww.com/NEN/A224). The nasal mucosa of 3 mice that had received long-term IN-I and 3 mice that
had received long-term IN-S did not have mucosal thickening
or inflammation.
Other attempts to have a more optimal comparison
group of diabetic mice receiving systemic insulin were
unsuccessful owing to infections resulting from repeated
glucose sampling during sliding scale approaches, placement
of venous catheters, and inefficacy of smaller doses of Subc
insulin (30).
Quantification of Insulin in Cerebrum and
Cervical Spinal Cord
Cerebral and spinal cord insulin concentrations were
greater at time points before STZ injection as well as in agematched nondiabetic mice versus diabetic mice receiving IN
saline (Table 1). When sampled at 1 hour after intervention,
D IN-I mice had higher insulin concentrations in brain and
spinal cord throughout 1 to 8 months of diabetes when compared with both D Subc-I and D IN-S mice (Table 1).
Quantification of Plasma Glucose After
Intranasal or Subcutaneous Intervention
In mice given Subc-IN, blood glucose levels fell over
hours; there was no significant decrease in mice receiving IN-I
or IN-S (Table 2). Unfortunately, this testing was performed
at an earlier time point of diabetes and did not capture the
impact of insulin-mediated hypoglycemia in C Subc-I and
D Subc-I mice recognized at older ages. Illness associated
with hypoglycemia and death over the subsequent week was
documented in 12 of 40 D Subc-I mice and 12/25 C Subc-I
mice between 4 and 9 months of age. We speculate that older
age was associated with greater likelihood of insulin-induced
hypoglycemia.
Impact of IN-I on Motor Behavioral Data
Before induction of diabetes, there were no baseline
differences in behavior testing between any mouse cohorts.
Overall rearing (supported and unsupported) was compromised in all diabetic cohorts but was better maintained in
the D IN-I group; over time, this group outperformed the D
Subc-I group (Fig. 2). Both individual hind limb and combined forelimb grip strengths were better preserved in D IN-I
mice over later stages of the experiments.
Impact of IN-I on Electrophysiology
Before induction of diabetes, there were no baseline
electrophysiological differences among the mouse cohorts.
Compound motor action potential amplitudes and MNCV
demonstrated age-related declines over time in nondiabetic
cohorts but with more rapid declines in diabetic cohorts
(Fig. 3A). Overall reductions in CMAP amplitudes and
MNCV began after 4 to 5 months of diabetes. In the later
months of diabetes (5Y7 months), D IN-I mice had better
maintained CMAPs and MNCV when compared with other
diabetic cohorts; there was only minimal protection from
electrophysiological changes in D Subc-I mice (Fig. 3B).
Estimated motor unit loss in diabetic mice was noticed
after 2 to 3 months of diabetes; a precipitous decline occurred
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J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
TABLE 3. Morphologic Features of Sciatic Nerves in
Nondiabetic (C) and Diabetic (D) Nerves From Mice Given
Intranasal or Subcutaneous Insulin or Saline After 8 Months
of Diabetes
Physical Property
Axonal fiber density (per mm2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Axonal area (Km2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Fascicular area (Km2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Myelination thickness (Km)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
G ratio
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
8 mo of Diabetes
15,024 T 235
14,793 T 241
14,918 T 263
14,714 T 221
14,622 T 369
14,606 T 376
14,607 T 291
14,598 T 302
38.3 T 0.5
37.7 T 0.6
38.2 T 0.6
37.9 T 0.4
34.9 T 0.7*†
31.7 T 0.6*
32.9 T 0.7*
31.6 T 0.6*
63.8 T 0.8
62.1 T 0.7
63.9 T 0.9
61.9 T 1.0
57.2 T 0.9*†
49.8 T 0.8*
53.8 T 1.0*
51.5 T 0.9*
1.08 T 0.04
1.03 T 0.03
1.09 T 0.04
1.02 T 0.04
0.97 T 0.03*‡
0.89 T 0.04*
0.94 T 0.05*
0.90 T 0.06*
0.60 T 0.03
0.61 T 0.03
0.60 T 0.04
0.61 T 0.03
0.61 T 0.04‡
0.63 T 0.04*
0.62 T 0.03*
0.63 T 0.04*
All measurements are mean T SEM.
G ratio was calculated as follows: axon area/(axon + myelin area).
*indicates significance at p G 0.016 with comparison to nondiabetic mice cohort
groups; †indicates significance with comparison to other diabetic cohort groups
(p G 0.016); and ‡indicates significance with comparison to Subc-S and IN-S diabetic
cohort groups using multiple ANOVA testing (p G 0.016) with Bonferroni post hoc t test
comparisons applied (> = 0.05).
IN-I, intranasal insulin; IN-S, intranasal saline; Subc-I, subcutaneous insulin; Subc-S,
delivery of subcutaneous saline.
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J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
TABLE 4. Morphologic Properties of Proximal and Distal
Peroneal Nerve Segments in Nondiabetic (C) and Diabetic (D)
Nerves From Mice Receiving Intranasal or Subcutaneous
Insulin or Saline After 5 or 8 Months of Diabetes
Physical Property
Proximal peroneal nerve
Axonal fiber density (per mm2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Axonal area (Km2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Fascicular area (Km2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Myelination thickness (Km)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
G ratio
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Distal peroneal nerve
Axonal fiber density (per mm2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
5 mo of Diabetes
8 mo of Diabetes
14,802 T 142
14,705 T 162
14,699 T 158
14,745 T 145
14,717 T 172
14,645 T 179
14,682 T 161
14,593 T 188
14,800 T 151
14,670 T 128
14,712 T 126
14,678 T 138
14,366 T 181*
13,912 T 151†
13,999 T 149†
13,688 T 164†
36.8 T 1.5
36.4 T 1.4
36.5 T 1.3
35.9 T 1.5
34.3 T 1.6
31.7 T 1.5
32.1 T 1.2
31.9 T 1.5
36.3 T 1.4
35.8 T 1.5
36.1 T 1.4
35.2 T 1.6
33.1 T 1.7*
29.6 T 1.4†
30.4 T 1.3†
29.1 T 1.4†
61.4 T 1.8
60.8 T 1.9
60.9 T 2.0
59.9 T 1.6
56.5 T 1.9*
51.7 T 2.1†
53.4 T 2.0†
50.7 T 2.0†
60.6 T 1.6
59.8 T 1.8
60.0 T 1.9
59.1 T 1.7
55.1 T 1.9*
49.2 T 2.0†
51.3 T 2.0†
48.4 T 1.9†
1.04 T 0.06
1.05 T 0.07
1.04 T 0.05
1.04 T 0.05
1.00 T 0.07
0.96 T 0.10
0.98 T 0.08
0.95 T 0.07
1.04 T 0.05
1.03 T 0.06
1.03 T 0.03
1.03 T 0.04
0.97 T 0.06
0.91 T 0.05†
0.94 T 0.06
0.89 T 0.05†
0.59 T 0.04
0.60 T 0.03
0.59 T 0.03
0.60 T 0.03
0.60 T 0.03
0.60 T 0.04
0.60 T 0.03
0.60 T 0.03
0.60 T 0.03
0.60 T 0.03
0.60 T 0.02
0.60 T 0.03
0.61 T 0.03
0.61 T 0.04
0.61 T 0.03
0.61 T 0.03
14,201 T
14,105 T
14,149 T
14,073 T
138
155
146
140
14,091 T 142
13,878 T 145
13,913 T 136
13,891 T 141
Motor Unit Loss in Diabetes
TABLE 4. (Continued)
Physical Property
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Axonal area (Km2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Fascicular area (Km2)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Myelination thickness (Km)
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
G ratio
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
5 mo of Diabetes
13,917 T 166
13,658 T 150†
13,782 T 151
13,553 T 148†
8 mo of Diabetes
13,512 T
12,821 T
13,006 T
12,911 T
130†*
160†
138†
152†
36.2 T 1.4
35.7 T 1.3
36.0 T 1.3
35.6 T 1.4
32.9 T 1.4
29.6 T 1.3†
30.1 T 1.4†
29.0 T 1.6†
35.7 T 1.5
35.1 T 1.4
36.0 T 1.5
34.8 T 1.3
30.9 T 1.4*†
26.1 T 1.5†
26.8 T 1.4†
25.8 T 1.7†
60.1 T 1.9
59.1 T 1.8
59.4 T 1.8
59.1 T 1.7
52.6 T 1.8*†
44.8 T 1.9†
47.2 T 2.0†
44.8 T 1.9†
59.4 T 1.8
58.2 T 1.7
59.3 T 1.7
58.0 T 1.6
50.3 T 1.9*†
41.5 T 1.8†
43.9 T 1.9†
41.3 T 2.0†
1.03 T 0.05
1.03 T 0.05
1.03 T 0.04
1.02 T 0.05
0.93 T 0.06
0.88 T 0.05†
0.90 T 0.06†
0.86 T 0.08†
1.02 T 0.04
1.01 T 0.07
1.02 T 0.05
1.01 T 0.06
0.93 T 0.05*
0.81 T 0.06†
0.85 T 0.05†
0.80 T 0.04†
0.59 T 0.03
0.60 T 0.03
0.60 T 0.03
0.60 T 0.03
0.61 T 0.04
0.61 T 0.04
0.61 T 0.04
0.62 T 0.03
0.59 T 0.03
0.60 T 0.02
0.60 T 0.03
0.60 T 0.03
0.61 T 0.03
0.62 T 0.04
0.61 T 0.03
0.62 T 0.03
All measurements are mean T SEM.
G ratio was calculated as follows: axon area/(axon + myelin area).
*indicates significance with comparison to other diabetic cohort groups (p G 0.016);
and †indicates significance at p G 0.016 with comparison to nondiabetic mice cohort
groups using multiple ANOVA testing with Bonferroni post hoc t test comparisons
applied (> = 0.05).
IN-I, intranasal insulin; IN-S, intranasal saline; Subc-I, subcutaneous insulin; Subc-S,
delivery of subcutaneous saline.
between 3 and 4 months of diabetes (Fig. 3C). Diabetic IN-I
mice had greater retention of motor units when compared
with other diabetic mouse cohorts at and after 5 to 6 months
of diabetes. For mice receiving Subc injections, D Subc-I
mice had greater numbers of motor units than D Subc-S mice
after 8 months of diabetes. Interestingly, C IN-I mice had
greater motor unit numbers than other nondiabetic cohorts
after 8 months.
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331
Francis et al
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J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Motor Unit Loss in Diabetes
TABLE 5. Forms of Terminal Innervation in Extensor Digitorum Brevis Muscles in Nondiabetic (C) and Diabetic (D) Mice
Receiving Intranasal or Subcutaneous Insulin or Saline After 8 Months of Diabetes
Extensor digitorum brevis muscle
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Tibialis anterior muscle
C IN-I mice (n = 6)
C IN-S mice (n = 6)
C Subc-I mice (n = 6)
C Subc-S mice (n = 6)
D IN-I mice (n = 5)
D IN-S mice (n = 4)
D Subc-I mice (n = 5)
D Subc-S mice (n = 5)
Preterminal Sprout (per mm3)
Ultraterminal Sprout (per mm3)
Nodal Sprout (per mm3)
13.4 T 2.5 (54%)
13.5 T 2.8 (58%)
13.3 T 2.1 (57%)
13 T 1.4 (59%)
12.2 T 1.6 (49%)
9.3 T 1.1* (51%)
10.2 T 1.2* (52%)
9.1T 1* (52%)
14.3 T 1.8 (27%)
12.6 T 1.6 (31%)
12.8 T 1.5 (30%)
13.2 T 1.9 (29%)
10.2 T 1.3 (25%)
8.0 T 1.4* (26%)
8.4 T 1.5* (24%)
8.1 T 1.2* (26%)
8.2 T 1 (19%)
6.9 T 0.9 (11%)
7.9 T 0.8 (13%)
7.4 T 1.1 (12%)
8.6 T 1.2 (26%)
7.9 T 1.4 (23%)
7.5 T 1.6 (24%)
6.8 T 1.4 (22%)
31.1 T 2.1 (52%)
30.2 T 1.8 (49%)
29.3 T 1.1 (51%)
31.0 T 1.5 (50%)
26.2 T 1.8*† (46%)
19.3 T 2.1* (48%)
19.8 T 2.4* (49%)
22.2 T 2* (48%)
16.8 T 1.3 (34%)
16.6 T 1.2 (35%)
14.8 T 1.1 (36%)
15.3 T 1.2 (35%)
13.2 T 1.3† (28%)
9.9 T 1.5* (28%)
9.1 T 1.2* (29%)
10.4 T 1* (31%)
9.6 T 1.1 (14%)
7.2 T 1 (16%)
9.4 T 1.9 (13%)
7.8 T 1.2 (15%)
9.6 T 1.3 (26%)
8.6 T 1.4 (24%)
8.2 T 1.2 (22%)
8.1 T 1.1 (21%)
All measurements are mean T SEM; numbers in parentheses are the percentages of the indicated sprout type for all sprouts.
Earlier time points of 1, 3, and 5 months did not show significant loss of sprout types in any diabetic groups. Similar loss of both preterminal and ultraterminal forms of sprouts was
seen with tibialis anterior muscle specimens from diabetic mice (not shown).
*p G 0.016 versus nondiabetic mice cohort groups; †indicates comparison to other diabetic cohort groups using multiple ANOVA testing with Bonferroni post hoc t test
comparisons (> = 0.05, p G 0.016).
IN-I, intranasal insulin; IN-S, intranasal saline; Subc-I, subcutaneous insulin; Subc-S, delivery of subcutaneous saline.
The initial SMUP amplitude increased in an agedependent manner for both nondiabetic and diabetic mice.
At and after 3 to 4 months, the initial SMUP was larger in
diabetic mice but was maintained in D IN-I mice (Fig. 3D).
Diabetic Subc-I mice were protected against initial SMUP
enlargement after 6 to 8 months of diabetes when compared
with D Subc-S mice.
Impact of IN-I on Peripheral Nerve Structure
There was axonal and fascicular atrophy present in the
sciatic nerves of diabetic mice without evidence of axonal loss
(Table 3). Diabetic sciatic nerve myelin thickness was also
reduced after 8 months of diabetes, with some protection
identified in D IN-I mice. G ratios were also preserved in the
sciatic nerves of D IN-I mice but not in other diabetic mice.
Peroneal nerves from diabetic mice were divided into a
proximal segment (section from branch of sciatic nerve 2 mm
in length) and a distal segment (2-mm section from proximal
to tibialis anterior innervation). The distal peroneal nerve
was subject to a loss of fiber density, axonal and fascicle
area (atrophy), and myelin thickness after 5 months of diabetes (Table 4, Figure, Supplemental Digital Content 2,
http://links.lww.com/NEN/A225), whereas similar changes
were detected in the more proximal peroneal nerve segment
only after 8 months of diabetes (Table 4). G ratios, however,
were unchanged for the peroneal nerves when diabetic and
nondiabetic cohorts were compared. There were no detected
pathological changes in diabetic mouse peroneal nerves after
1 and 3 months of diabetes (data not shown). Diabetic IN-I
mice were protected from declines in distal peroneal nerve
axon density and from axonal atrophy as well as loss of
myelin thickness in both the proximal and the distal peroneal
nerve segments after 8 months of diabetes when compared
with diabetic Subc-I, Subc-S, and IN-S mice.
FIGURE 4. Motor end plate innervations in distal leg muscles from mice with diabetes [D] and without [C]. Delivery of subcutaneous saline is indicated as ‘‘Subc-S,’’ subcutaneous insulin as ‘‘Subc-I,’’ intranasal saline as ‘‘IN-S,’’ and intranasal insulin as ‘‘IN-I.’’
(AYC) Selected confocal images are demonstrated for the tibialis anterior muscle from a C IN-I mouse (A), D IN-I mouse (B), and D
IN-S mouse (C). Axons are identified with immunohistochemistry for NF-200 (green); neuromuscular junctions (NMJs) are demonstrated using >-bungarotoxin (BG). White thick arrows indicate terminal NMJ innervation; white thin arrows, axons forming
preterminal sprouts; arrowheads, NMJs failing to receive innervation. (DYF) Examples of forms of sprouting are shown from
nondiabetic mice samples, including ultraterminal sprouting (D), preterminal sprouting (E), and nodal sprouting (F). Bar = 10 Km
for AYC and 4 Km for DYF. (GYJ) Diabetes was associated with the loss of NMJs (G, I) and end plate terminal innervation (H, J),
with partial preservation in D IN-I mice (see also Table 5). Significant differences were determined by multiple repeated-measures
ANOVA tests, with *indicating significant difference (> G 0.05, p G 0.016 using Bonferroni corrections) between the D IN-I mouse
group and other diabetic mouse cohorts.
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333
Francis et al
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Impact of IN-I on Motor Neurons
and Innervation
Motor neurons within lamina IX of the lumbar
spinal cord were unchanged in neuronal density or neuronal
area in any diabetic or nondiabetic mouse cohorts at 1, 3,
5, or 8 months (Figure, Supplemental Digital Content 2,
http://links.lww.com/NEN/A225 and data not shown). There
were no differences in the morphology (neuronal area, presence of vacuolation) of motor neurons in the corresponding
cervical or thoracic regions of the spinal cord (not shown).
Vacuolation was rarely identified in any cohort.
All mice cohorts demonstrated age-related decline in
numbers of NMJs and amounts of terminal end plate innervation over time (Fig. 4 and Table 5). Diabetic mice had
further acceleration of loss of both NMJs and terminal end
plate innervation (Fig. 4) within 2 distal muscles: extensor
digitorum brevis and tibialis anterior. After 8 months of diabetes, both preterminal and ultraterminal forms of sprouts were
reduced, but there was no loss of nodal sprouts (Table 5).
Diabetic IN-I mice, however, were protected from loss of preterminal and ultraterminal axonal sprouts as well as from loss
of overall NMJs and terminal end plate innervation in both
distal muscles (Fig. 4). Several diabetic mice specimens had
terminal nerve endings with end bulbs identified, but end bulbs
were not quantified.
Most extensor digitorum brevis sections examined for
fiber-type morphology were unremarkable in both diabetic
and nondiabetic mice. However, a few regions of the extensor digitorum brevis from diabetic mice demonstrated some
histologic changes typical of neurogenic denervation after
8 months of diabetes (Figure, Supplemental Digital Content 3,
http://links.lww.com/NEN/A226).
Impact of IN-I on Signaling Pathways in
Diabetic Spinal Cord
Lumbar spinal cord was obtained at 1, 2, 4, 8, and
24 hours after IN and Subc delivery of insulin or saline
to measure proteins and ratios of phosphorylated-total
protein in the PI3K/Akt pathway in diabetic and nondiabetic mice (Figure, Supplemental Digital Content 4,
http://links.lww.com/NEN/A227). Intranasal delivery of insulin led to increased phosphorylationYtotal Akt, GSK-3A, and
CREB at 2 hours, with loss of phosphorylated-total CREB after
24 hours. Subcutaneous insulin led to rises in phosphorylatedtotal Akt, GSK-3A, and CREB at 8 hours after Subc-I, with
subsequent declines at 24 hours. Intranasal delivery of insulin
for 2 days only after 8 months of untreated diabetes led to
short-term rises in phosphorylation for each ERK, MEK, and
JNK not identified after 7 days of IN-I delivery at the end of
8 months of diabetes. Long-term (8 months) IN-I delivery
during the entire duration of diabetes also did not lead to rises
in phosphorylated MEK, ERK, or JNK (Figure, Supplemental
Digital Content 5, http://links.lww.com/NEN/A228).
Diabetes of 8 months in duration was associated with
a loss of IR-mediated signaling molecules within the spinal
cord, particularly in the lumbar spinal region (Fig. 5A).
Quantification analysis of protein blots in the lumbar cord
region identified a loss of each of PI3K, pAkt, Akt, pGSK-3A,
334
FIGURE 5. After 8 months of diabetes, Western blotting identified generalized diminution of protein expression within diabetic
spinal cord regions compared with nondiabetic spinal regions,
except for NF-JB. Delivery of subcutaneous saline is indicated
as ‘‘Subc-S,’’ subcutaneous insulin as ‘‘Subc-I,’’ intranasal saline
as ‘‘IN-S,’’ and intranasal insulin as ‘‘IN-I.’’ (A) A sample protein
blot for downstream molecules related to insulin signaling
(PI3K, pAKT, Akt, pGSK-3A, GSK-3A, pCREB, CREB, IRS-1, and
IRA), and the transcription factor NF-JB. A-Actin was used as a
loading control and for relative quantification in all cases. (B)
Semiquantification of 3 individual Western blots for each mouse
cohort identified downregulation in each for PI3K and the
ratios of pAkt/Akt, pGSK-3A/GSK-3A, and pCREB/CREB for
the lumbar spinal cord region. Multiple ANOVA tests were performed in each case; *indicates significant difference (> G 0.05,
p G 0.016) between D IN-I and D Subc-I cohorts; ?, significant
difference (> G 0.05, p G 0.016) between D IN-I and D IN-S
cohorts. Only data for C Subc-S, D Subc-S, D Subc-I, D IN-S, and
D IN-I are shown.
GSK-3A, pCREB, and CREB for all diabetic lumbar cord
tissue (Fig. 5B). However, protection against relative loss of
pAkt, pGSK-3A, and pCREB occurred in D IN-I mice. No
consistent changes could be identified at the level of the sciatic or peroneal nerve for these insulin-associated markers.
Diabetes was also associated with a relative loss of
intraneuronal pAkt and decreased nuclear pAkt presence
in spinal motor neurons using immunohistochemical measures (Fig. 6), with approximately 40% more nuclear pAkt
positivity identified in D Subc-I mice and 90% more seen in D
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J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Motor Unit Loss in Diabetes
FIGURE 6. IR-A and pAkt expression within spinal motor neurons. Delivery of subcutaneous saline is indicated as ‘‘Subc-S,’’ subcutaneous insulin as ‘‘Subc-I,’’ intranasal saline as ‘‘IN-S,’’ and intranasal insulin as ‘‘IN-I.’’ (AYC) Colocalization is demonstrated for
C IN-I mice (A), D IN-I mice (B) and D IN-S mice (C). (D, E) Quantitative real-time polymerase chain reaction (qRT-PCR) identified
marked downregulation for PI3K (D) and Akt (E) mRNA in spinal cord from diabetic mice; the greatest loss was in the lumbar
regions. Diabetic IN-I mice had protection against PI3K and Akt loss in spinal cord, greatest in the lumbar regions. Multiple ANOVA
tests were performed, with *indicating significant difference (> G 0.05, p G 0.16) between groups indicated by horizontal bars.
IN-I mice. Among nondiabetic mouse cohorts, there was no
significant difference in pAkt protein presence. Quantification of mRNA and protein for PI3K and Akt demonstrated
downregulation with diabetes in general, with partial protection from downregulation for both PI3K and Akt mRNA
identified in D IN-I mice in lumbar and thoracic spinal cord;
protection against PI3K downregulation was only seen in
cervical spinal cord in D IN-I mice. No consistent changes
in mRNA for PI3K signaling pathway molecules could be
identified in the sciatic or peroneal nerves of diabetic mice
receiving intervention.
DISCUSSION
Intranasal insulin protected against the loss of motor
function and distal motor end-terminals in diabetic mice; it
slowed the development of estimated motor unit loss, distal
axonal retraction, and distal axonal atrophy and loss. Intranasal delivery of insulin, moreover, avoids the systemic adverse effects associated with Subc-I therapy in this model.
We hypothesize that insulin likely exerts its beneficial effects
through the PI3K/Akt signaling pathway downstream of the
IR. The benefits of IN-I in the diabetic nervous system dem-
onstrated parallel those benefits on the sensory neuron and
brain in the diabetic nervous system (29, 30).
Although changes in the motor unit develop later than
those of sensory neurons and their axons in diabetes (7),
they are also subject to neurodegeneration (6, 8). Alterations
in diabetic motor neurons begin first at the distal-most end
plate terminals with a loss of end plate innervation, followed
by loss of NMJs, development of axonal atrophy, and even
axonal loss within distal portions of motor-dominant peripheral nerves. There is a mild age-dependent loss of both
CMAP amplitude and MNCV detectable over time, which is
not evident at earlier time points (10) but is exacerbated for
at least 2 to 3 months of diabetes exposure (7). Despite this,
there are no detectable morphologic changes occurring at the
motor neuron perikarya after marked behavioral and electrophysiological changes have already been detected.
The STZ-induced mouse model of diabetes has the advantages of high rates of diabetic conversion and prolonged
life span. Here, this mouse model provided studies of sufficient length to capture changes only present after several
months of diabetes, whereas shorter-term models would be
unlikely to identify such abnormalities. The loss of motor end
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335
Francis et al
J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
plates is likely the consequence of distal retraction of axon
terminals (34, 35). Previously identified ultrastructural changes
in the NMJ of the diabetic mouse (6) are similar to those
found with retraction of terminals in motor neuron disorders.
In diabetic murine muscle samples, ultrastructural changes in
the NMJ accompany reduced acetylcholine receptor expression (36). We hypothesize that overall retraction of nerve endterminals develops progressively while advancing proximally;
motor neuronal perikaryal structural changes and drop out, if
these occur, are the final events in this process of motor unit
loss related to diabetes.
sulin signaling without promoting hypoglycemia, can also
promote gene responses in the diabetic peripheral nervous
system (41). This strategy may be beneficial in humans with
diabetic neuropathy (42). In motor neurons, insulin stimulation may promote the maintenance of distal end plate terminal
innervation. Insulin upregulates proteinYtyrosine phosphorylation (43) through downstream activation of IRS-2 (44), thereby
promoting the activation of Akt and phosphorylation of Akt
substrates (45). Akt activation is also important for motor
neuron survival after sciatic nerve injury (46, 47), possibly
owing to the activation of NF-JB or the inhibition of proapoptotic factors (48). In our studies, insulin-mediated activation of the PI3K/Akt pathway (44, 49) led to phosphorylation
of Akt effectors including CREB and GSK-3A (50Y53).
Phosphorylation of CREB inhibits apoptosis in neurons (54),
whereas the loss of CREB results in impaired axonal growth
(55). We have found that IN-I was associated with elevation
of pCREB levels (56) and pGSK-3A (56) within diabetic
mouse spinal cord. Conversely, long-term insulin exposure
failed to maintain long-term activation of the MEK/ERK and
JNK pathways, and short-term exposure transiently seemed
to reverse phosphorylation of these alternative insulin signaling pathways (Figure, Supplemental Digital Content 5,
http://links.lww.com/NEN/A228). These results indicate that
insulin’s actions are at least partially mediated through the
PI3K/Akt pathway within CNS motor neurons.
Neural and Systemic Effects of Subc-I and IN-I
We previously quantified IN and Subc radiolabeled insulin delivery and determined that peak delivery to the brain
and spinal cord occurs within 1 hour after IN delivery and
Subc delivery peaks approximately 6 hours later (29, 30).
Insulin concentrations at the thoracic and lumbar spinal cord
regions were similar between IN and Subc insulin delivery.
Whereas mice receiving IN insulin maintained good health
throughout the 1-, 2-, and 6-hour monitoring periods before
death, mice receiving Subc insulin had much higher concentrations of insulin in whole blood and nonnervous organs;
nearly half of these mice developed hypoglycemia-induced
illness (29, 30).
The 2 forms of insulin delivery had important differences on outcomes. Subcutaneous delivery of insulin was
inferior to IN-I in diabetic mice owing to the greater mortality related to episodes of hypoglycemia. In addition, Subc-I
led to improved glycated hemoglobin A1c levels (unlike in D
IN-I mice), suggesting that the beneficial effects of IN-I
in diabetes are not primarily related to corrections in hyperglycemia; this may not be true with Subc-I. Intranasal delivery of insulin to the cortex and brainstem (where central
motor neurons reside) was also greater and more rapid than
with Subc-I, possibly contributing to the protection of central
diabetes-mediated neurodegeneration (29).
Insulin as a Neuroprotective Factor and
Downstream Signaling Pathways
The main site of insulin activity, the IR, is present on
motor neurons in the spinal cord, central glial cells, on myelinated anterior root fibers, and in the spinal cord (23, 37).
Although intrathecal insulin prevents degeneration and promotes regeneration in injured peripheral nerve (23), the
greatest effect of insulin may be the prevention of a ‘‘dyingback’’ pathophysiological process beginning in the most
distal epidermal fibers exposed to diabetes (38). Our results
indicate that a similar distal-first process occurs within
motor pathways in the spinal cord and peripheral nervous
system. Although motor neuron loss could not be detected, it
is possible that longer-duration models with sustaining
therapies will lead to recognition of morphologic changes in
motor neurons. Indeed, humans with diabetes can develop
spinal cord atrophy (39, 40).
Insulin stimulation of the IR and its subsequent signaling pathways protects neurons from the effects of diabetes
through multiple mechanisms (25). Previous studies have
demonstrated that proinsulin C-peptide, which stimulates in-
336
Insulin’s Role at the Motor Unit in Diabetes
Motor unit recruitment is based on Henneman’s size
principle, with smaller motor units firing before larger motor
units (57). Among diabetic mice, the initial SMUP amplitude
increased, suggesting that a preferential loss of smaller motor
units or their enlargement occurred owing to a compensatory
sprouting to denervated muscle fibers (9). Diabetic mice had
fewer overall preterminal and ultraterminal sprouts when
compared with nondiabetic mice, as well as less terminal end
plate innervation. The terminal forms and nodal sprouts were
higher in diabetic mice when compared with nondiabetic
mice, possibly because of the attempts to reinnervate muscle fibers that have been denervated, leading to motor unit
enlargement. Insulin may promote such compensatory axonal
sprouting (58). It remains possible that some of the effects of
insulin may be cerebral in origin, maintaining cerebrospinal
motor pathways; the early benefit on rearing studies may
support this theory. It is unlikely that IN-I is unlikely to gain
access to the peripheral nerve to influence insulin-deficient
Schwann cells, perhaps leading to less consistent benefits in
MNCV testing. It is also possible that Subc-I levels needed
to support potentially insulin-deficient Schwann cells cannot
be achieved without the occurrence of problematic systemic
hypoglycemia.
Accompanying the lost innervation and increased initial SMUP amplitude was a loss of motor units estimated by
MUNE. It has been demonstrated that insulin signaling plays
a prominent role in downstream function, including synaptic
plasticity and density (59). Similar functions may permit insulin to act at the motor neuron, preventing downstream loss of
denervation. Similarly, insulin also plays a role at the NMJ. It
has been suggested that short-range diffusible sprout-inducing
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J Neuropathol Exp Neurol Volume 70, Number 5, May 2011
Motor Unit Loss in Diabetes
factors may be released from denervated or inactive muscle
fibers, including the insulin-like growth factors (60). This may
be related to the developmental presence of insulin-like receptors and insulin-like peptides identified in other organisms
(61). In the present study, the superiority of IN insulin over
systemically delivered insulin suggests that the more potent
effect is at the motor neuron.
testing and rearing is uncertain but is likely substantial, potentially confounding the motor testing results obtained. Our
calculated MUNE values are higher than those reported in
the literature for both rodents (10, 66) and humans (67); this
may relate to the user-dependent technique of incremental
MUNE methods, problems with comparing different methods
of MUNE, differences in muscles studied, and differences in
mouse strains. We have also not confirmed that the mode of
action of insulin is restricted to the PI3K/Akt signaling pathway; other substrates and signaling pathways such as with
the RasYmitogen-activated protein kinase pathway, which
regulates the expression of genes for cell growth and differentiation through activation of ERK/MEK (68), Rac, CDC42,
and JNK 1Y3 (69, 70), may be important. Moreover, the impact of insulin on other substrates such as Gab-1, p60dok,
Cbl, APS, Shc isoforms, phosphotyrosine phosphatase SHP2,
and the cytoplasmic tyrosine kinase Fyn (71) may also be important at the motor system. Based on the present results,
however, it did not seem as though ERK/MEK is pertinent
to the changes described (Figure, Supplemental Digital Content 5, http://links.lww.com/NEN/A228). Finally, we have not
shown evidence of a cause and effect for IN insulin delivery
and the enhanced phosphorylation of proteins in the PI3K/Akt
pathway; this remains associative at present. Nevertheless, IN-I
seems to be a prospective, potent therapy for the amelioration
of the behavioral, electrophysiological, morphologic, and molecular changes that occur in the diabetic motor unit. Its actions seem to be independent of its effects on glycemia, and
our results support its role as an important trophic factor in the
management of diabetic nervous system complications.
Utility of Intranasal Delivery in Patients With
Diabetic Nervous System Complications
In situations where insulin deficiency contributes to
nervous system dysfunction, raising systemic insulin levels is
unlikely to be an effective long-term strategy owing to the
risks of hypoglycemia. Intranasal administration bypasses
the blood-brain barrier, targeting insulin to the brain, spinal
cord, and cerebrospinal fluid within 1 hour (29, 30). Extracellular bulk flow transport along both olfactory and trigeminal neural pathways likely carries IN-I, possibly using
perivascular channels of blood vessels entering the CNS (28).
In humans, plasma glucose levels may be mildly decreased
by IN insulin delivery, but patients remain euglycemic (62).
Although the use of IN insulin for the management of systemic diabetes has been limited to date, in our mouse cohorts,
D IN-I mice had better maintenance of body weight and less
mortality. The reason for this is unclear but may relate to less
severe sensorimotor decline and cognition loss; such ailments
may contribute to weight loss in Alzheimer disease (63) and
motor neuron disease patients (64). Although IN-I did not
alter blood glucose levels, Subc-I diminished hyperglycemia,
making it difficult to separate the relative contributions of
antihyperglycemic actions from the trophic properties of insulin with Subc-I (65).
We acknowledge some limitations of our present
results. First, although changes in the human motor system
occur with diabetes, these occur at late stages of DPN and are
modest in comparison to the sensory manifestations. These
results must be considered under the limitations of inability
to achieve a more appropriate long-term diabetic control
cohort with optimal glycemic management (29, 30). Previous
attempts to use either glycemia-regulated dosing or lesser
amounts of Subc insulin led to unacceptable levels of mortality or inadequate and unpredictable results in glycemic
levels (29, 30). Hypoglycemia may have affected some of
the results of motor testing; the impact of hypoglycemia on
the D Subc-I and C Subc-I cohort groups was anticipated,
but unavoidable. Indeed, mortality in the C Subc-I mice was
greater than that of the C Subc-S mice, indicating that hypoglycemia induced by insulin was problematic for mice of the
Subc cohort. The form of insulin used (Humulin R) was shortacting and was selected based on prior experience (23, 24),
but use of a longer-acting form of insulin may have been more
optimal. Future studies using Subc insulin pellets may be
more advantageous for the preservation of long-term diabetic
models compared with mice receiving IN insulin. Although
protection against motor unit loss, distal axonal retraction,
atrophy, and loss was identified, some parameters such as
motor conduction velocities failed to demonstrate consistent
benefits with IN-I. The impact of sensory loss and neuropathic
pain behaviors on motor testing such as with grip strength
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