Content uploaded by Hans-Christer Holmberg
Author content
All content in this area was uploaded by Hans-Christer Holmberg on Jul 19, 2018
Content may be subject to copyright.
The Journal of Physiology
J Physiol 589.3 (2011) pp 711–725 711
Role of glycogen availability in sarcoplasmic reticulum
Ca2+kinetics in human skeletal muscle
Niels Ørtenblad1,JoachimNielsen
1,BengtSaltin
2and Hans-Christer Holmberg3,4
1University of Southern Denmark, Odense, Denmark
2Copenhagen Muscle Research Centre, Copenhagen, Denmark
3Swedish Winter Sports Research Centre, Department of Health Sciences, Mid Sweden University, Sweden
4Swedish Olympic Committee, Stockholm, Sweden
Non-technical summary Glucose is stored as glycogen in skeletal muscle. The importance of
glycogen as a fuel during exercise has been recognized since the 1960s; however, little is known
about the precise mechanism that relates skeletal muscle glycogen to muscle fatigue. We show that
low muscle glycogen is associated with an impairment of muscle ability to release Ca2+,which
is an important signal in the muscle activation. Thus, depletion of glycogen during prolonged,
exhausting exercise may contribute to muscle fatigue by causing decreased Ca2+release inside
the muscle. These data provide indications of a signal that links energy utilization, i.e. muscle
contraction, with the energy content in the muscle, thereby inhibiting a detrimental depletion of
the muscle energy store.
Abstract Little is known about the precise mechanism that relates skeletal muscle glycogen to
muscle fatigue. The aim of the present study was to examine the effect of glycogen on sarcoplasmic
reticulum (SR) function in the arm and leg muscles of elite cross-country skiers (n=10, ˙
VO2max
72 ±2mlkg
−1min−1) before, immediately after, and 4 h and 22 h after a fatiguing 1 h ski race.
During the first 4 h recovery, skiers received either water or carbohydrate (CHO) and thereafter all
received CHO-enriched food. Immediately after the race, arm glycogen was reduced to 31 ±4%
and SR Ca2+release rate decreased to 85 ±2% of initial levels. Glycogen noticeably recovered after
4 h recovery with CHO (59 ±5% initial) and the SR Ca2+release rate returned to pre-exercise
levels. However, in the absence of CHO during the first 4 h recovery, glycogen and the SR Ca2+
release rate remained unchanged (29 ±2% and 77 ±8%, respectively), with both parameters
becoming normal after the remaining 18 h recovery with CHO. Leg muscle glycogen decreased to
a lesser extent (71 ±10% initial), with no effects on the SR Ca2+release rate. Interestingly, trans-
mission electron microscopy (TEM) analysis revealed that the specific pool of intramyofibrillar
glycogen, representing 10–15% of total glycogen, was highly significantly correlated with the SR
Ca2+release rate. These observations strongly indicate that low glycogen and especially intra-
myofibrillar glycogen, as suggested by TEM, modulate the SR Ca2+release rate in highly trained
subjects. Thus, low glycogen during exercise may contribute to fatigue by causing a decreased SR
Ca2+release rate.
(Received 6 July 2010, accepted after revision 29 November 2010; first published online 6 December 2010)
Corresponding author N. Ørtenblad: Institute of Sports Science and Clinical Biomechanics, University of Southern
Denmark,Campusvej 55, 5230 Odense M, Denmark. Email: nortenblad@health.sdu.dk
Abbreviations cc, cross country; CHO, carbohydrate; E–C, excitation–contraction; HR, heart rate; HRmax, maximum
heart rate; [La−], lactate concentration; IMF, intermyofibrillar; Intra, intramyofibrillar; MHC, myosin heavy chain; PCr,
phosphocreatine; RyR; ryanodine receptor (Ca2+release channel); SR, sarcoplasmic reticulum; SS, subsarcolemmal;
TEM, transmission electron microscopy.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society DOI: 10.1113/jphysiol.2010.195982
712 N. Ørtenblad and others J Physiol 589.3
Introduction
In skeletal muscle, glucose is stored as glycogen, which is
a major source of energy during most forms of muscle
activity. Studies at the beginning of the last century by
Krogh and Lindhard (Krogh & Lindhard, 1920), and
later by Christensen and Hansen (Christensen & Hansen,
1939), revealed the importance of carbohydrate as a
fuel during exercise. Later the use of the muscle biopsy
technique demonstrated a direct correlation between
muscle glycogen concentration and time to fatigue during
moderately intense exercise (∼75% of maximum oxygen
uptake; ˙
VO2max)(Bergstr¨
om et al. 1967). The importance of
muscle glycogen on performance during both prolonged
and high intensity intermittent exercise has subsequently
been confirmed in numerous studies (for a review
see Green, 1991). However, the link between glycogen
depletion and the development of fatigue, as well as the
precise mechanism whereby muscle glycogen affects the
series of events that ultimately result in fatigue, is not fully
understood.
The existence of a glycogenolytic complex associated
with sarcoplasmic reticulum (SR) is now well established
(Wanson & Drochmans, 1972; Entman et al. 1980;
Xu & Becker, 1998). This SR–glycogen complex
consists of glycogen and associated proteins involved in
glycogenolysis and resynthesis, glycolysis and regulating
proteins. These assemblies are dynamic and raise the
possibility that feedback regulation from glycogen storage
deposits associated with the SR may exist. Such an
SR–glycogen arrangement may permit the regulation of
the level of [Ca2+]iand, in turn, the regulation of the
contractile activation and energy utilization of skeletal
muscle.
Several investigators have demonstrated impaired SR
function following exercise (Byrd et al. 1989; Favero et al.
1993; Ørtenblad et al. 2000; Duhamel et al. 2006a). These
data are obtained in homogenates from fatigued muscle by
direct measurements of SR vesicle Ca2+-ATPase activity,
and SR vesicle uptake and release rates. The idea of an
exercise-induced impaired SR function, leading to muscle
fatigue, is supported by experiments on single fibres with
simultaneous measurements of force and [Ca2+]i.These
experiments have demonstrated that [Ca2+]iis decreased
during repeated tetani, which is primarily caused by
a decreased SR Ca2+release rate (Allen et al. 2008).
While it is generally accepted that impaired SR Ca2+
release is a substantial factor in fatigue following most
forms of exercise, several mechanism can potentially be
involved in the series of events that lead to a decrease
in SR Ca2+release rate. A range of different studies
suggests that reduced muscle glycogen results in an
impairment of SR function. Using both single fibres and
muscle bundles, Chin and Allen (Chin & Allen, 1997)
elegantly demonstrated, through the manipulation of
glucose availability in the recovery phase after fatiguing
contractions, that muscle force and [Ca2+]iare associated
with muscle glycogen content. Thus, a reduced resting level
of glycogen was associated with a faster decrease in tetanic
[Ca2+]iand force during subsequent contractions. These
results have subsequently been confirmed (Kabbara et al.
2000; Helander et al. 2002) and together these data suggest
that the change in SR function associated with fatigue and
recovery has a component which is glycogen dependent.
Additional evidence for a link between muscle glycogen
and SR function comes from human experiments in
which Duhamel and colleagues manipulated pre-exercise
glycogen in untrained subjects and thereby demonstrated
an association between low muscle glycogen levels and a
reduced SR vesicle Ca2+release rate during subsequent
exercise (Duhamel et al. 2006b,c). In line with this, by
using mechanically skinned fibres from toad and rat EDL,
where ATP and phosphocreatine (PCr) levels can be kept
high and constant, muscle glycogen content has been
correlated with the muscle fatigability (Stephenson et al.
1999; Barnes et al. 2001; Nielsen et al. 2009). Thus, despite
a constant myoplasmic ATP and PCr concentration,
muscle glycogen level affects the excitation–contraction
(E–C) coupling. This is interpreted as glycogen having
either a non-metabolic role or that compartmentalized
energy transfer through the glycolysis is important for
maintaining normal E–C coupling (Chin & Allen, 1997;
Stephenson et al. 1999; Barnes et al. 2001; Nielsen et al.
2009).
In agreement with the hypothesis of a role for
glycogen in E–C coupling, transmission electron
microscopy (TEM) reveals that glycogen is located in
distinct compartments close to different sites of E–C
coupling and with possible distinct functions (Wanson &
Drochmans, 1972; Sj¨
ostr¨
om et al. 1982a;Fridenet al. 1985,
1989; Marchand et al. 2002, 2007). Thus, muscle glycogen
is located: (1) in the subsarcolemmal region beneath the
sarcolemma (SS), (2) between the myofibrils close to SR
and mitochondria (IMF), and (3) inside the myofibrils,
between the contractile filaments, mainly in the I-band
(Intra). In mechanically skinned fibres, we have previously
shown distinct functions for these subpopulations in rat
muscle, with the intramyofibrillar glycogen correlating
with fatigue resistance capacity, and the intermyofibrillar
glycogen correlating with the muscle half-relaxation time
in an unfatigued tetanic contraction (Nielsen et al.
2009). Furthermore, it is recognized that repeated contra-
ctions mediate an uneven breakdown of the three sub-
populations of glycogen (Sj¨
ostr¨
om et al. 1982b;Friden
et al. 1985, 1989; Marchand et al. 2007). However,
while there is evidence for an association between low
muscle glycogen and SR function, little is known about
the role of the individual glycogen compartments and
SR function. Furthermore, the association between low
muscle glycogen and exhaustion is less pronounced in
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 713
highly trained subjects and it remains uncertain whether
there is an association between muscle glycogen and SR
function in these athletes. In the present study we have
measured muscle glycogen content and the SR function
in humans in the recovery period after glycogen-depleting
exercise. Thus, by either keeping muscle glycogen low or
allowing optimal glycogen resynthesis during recovery, we
were able to create an experimental design where changes
in muscle glycogen level were separated from parallel
changes affecting SR Ca2+function.
The aim of the present study was to investigate the
role of skeletal muscle glycogen on SR function in elite
cross country (cc) skiers, through manipulating glycogen
levels with exhaustive arm and leg exercise and variation
in the CHO feeding in the recovery after the exercise.
We hypothesize that low glycogen levels are associated
with impaired SR function and that restricting CHO
intake in the early recovery period after exhaustive exercise
will hinder the recovery of SR function. Furthermore, by
using electron microscopy images to visualize and quantify
muscle glycogen granules, we aimed to examine the role of
possible distinct functions of compartmentalized glycogen
on the SR function in trained humans. Thus, the specific
hypothesis to be tested was that low intramyofibrillar
glycogen content, localized near the SR Ca2+release site,
is associated with SR Ca2+release and intermyofibrillar
glycogen is related to the SR Ca2+uptake rate.
Methods
Ethical approval
The project was approved by the Regional Ethical Review
BoardinUme
˚
a, Sweden (no. 07-076M). Before giving
their written consent to participate, the subjects were fully
informed about the project, the risks involved, discomfort
associated with the experiment, and that they could
withdraw from the project at any time.
Subjects
Ten male elite cc-skiers (age: (mean ±S.E.M.)
22 ±0.4 years; height: 181 ±2 cm; body mass:
78.8 ±2.6 kg) with a ˙
VO2maxof 72 ±2 (range 62–79)
ml kg−1min−1(5.4 ±0.5 (range 4.8–6.1) l min−1)and
haemoglobin concentration of 155 ±2gl
−1, volunteered
for the study. The skiers were selected from Norwegian
elite skiers and had an average of 11 years of training. Six
out of ten competed in the Norwegian national team and
eight competed in the FIS World Cup the subsequent year;
one subject had won a world cup race. All participants
were training an average of 700 h per year (550–850 h).
Experimental design
In order to validate the role of muscle glycogen levels in SR
function the cc-skiers completed a high intensity classical
cc race, reducing muscle glycogen levels in both arms and
legs, and were then followed in the recovery period after
the race. During the first 4 h recovery, the skiers were
randomized and received either water or carbohydrate
(CHO), which allowed discrimination between muscle
glycogen level without having an effect of acute exercise
on SR function. After the 4 h recovery, the subject received
the same CHO-enriched energy intake for the remainder
of the 22 h recovery, i.e. 18 h CHO for the water trial.
The CHO-enriched energy intake consisted of both liquid
solutions and food allowing the skiers to receive at least 1 g
CHO kg−1h−1. Muscle biopsies were obtained in both arm
and leg before (Pre), immediately after (Post), as well as 4 h
and 22 h after the cc race. The Post biopsy was obtained
immediately after (1–2 min) the skiers had crossed the
finish line. Heart rate (HR) was measured continuously
during the race and blood lactate concentration ([La−])
was measured 1 min after the finish. The cc-skiers were
tested for ˙
VO2max in the laboratory within 2 weeks of the
performance test.
Performance
Each subject performed a ∼20 km time trial (classic style)
on a competition (hilly) cc racecourse. The race and tests
took place at the end of the race season. In order to ensure
maximum performance, the time trial was arranged as
a competition between the participating subjects, with
the chance to win bonus money for the three best final
times and time at half-distance. The weather conditions
during the ski race were optimal with a clear sky, no wind,
an air and snow temperature of ∼−2◦Candarelative
humidity of 70%. Subjects performed individual warm-up
and ski preparation before the race. Snow conditions and
the racecourse favoured the diagonal stride technique and
a high degree of double poling, i.e. upper body exercise.
Laboratory tests
˙
VO2max and maximal heart rate (HRmax) were determined
using diagonal skiing on a treadmill (Rodby, S¨
odert¨
alje,
Sweden) (Calbet et al. 2005). The skiers started
at 11 km h−1on a treadmill inclination of 4 deg;
subsequently the inclination was raised 1 deg every minute
to exhaustion. Blood samples (20 μl) were taken from the
fingertip and used for determination of [La−]byBiosen
5140 (EKF-diagnostic GmbH, Magdeburg, Germany). All
tests were performed using roller skis on a motor-driven
treadmill (Rodby RL 3000, V¨
ange, Sweden), secured in a
safety harness suspended from the ceiling. All participants
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
714 N. Ørtenblad and others J Physiol 589.3
were well accustomed to roller skiing on the treadmill as
part of their regular training.
Respiratory variables were measured with the mixed
expired procedure employing an ergo-spirometry system
(AMIS 2001 model C, Innovision A/S, Odense, Denmark),
equipped with an inspiratory flowmeter. The gas analysers
were calibrated with a high-precision two-component
gas mixture of 16.0% O2and 4.0% CO2(Air Liquide,
Kungs¨
angen, Sweden) and calibration of the flowmeter
was performed at low, medium and high flow rates
with a 3-l air syringe (Hans Rudolph, Kansas City,
MO, USA). Ambient conditions were monitored with an
external apparatus (Vaisala PTU 200, Vaisala OY, Helsinki,
Finland). Expired O2and CO2and the inspired minute
ventilation ( ˙
VE) were monitored continuously and ˙
VO2
values were calculated and averaged during the final 30 s
at each workload. HR was followed continuously using
the HR monitor Polar S610 (Polar Electro OY, Kempele,
Finland) in combination with the metabolic cart.
Muscle biopsy preparation and analytical procedures
Muscle biopsies were obtained in both arm and leg muscles
before and immediately after the race (within 1–2 min after
the race), as well as 4 h and 22 h after the race. The biopsies
weretakeninarandomizedorderontheleftandright
side, with two biopsies in each arm and leg. All biopsies
from individual subjects were taken by the same person to
ensure standardization of the location on the muscle and
muscle depth from where the biopsies were taken. After
local anaesthesia (2–3 ml 2% lidocaine (lignocaine)), an
incision was made through the skin and fascia and the
muscle biopsy was taken from m. vastus lateralis (leg)
and m. triceps brachii (distal part of the lateral head,
arm), using a modified Bergstr¨
om needle with suction.
These muscles were preferred because they are highly
active during cc skiing (Holmberg et al. 2005). The muscle
specimenwasdriedonafilterpaperplacedonaglass
plate cooled on ice. After the removal of visible connective
tissue and fat the muscle specimen was divided into three
specimens and either: (1) frozen directly in liquid N2
and stored for later analyses of metabolites, (2) a muscle
segment was fixed for transmission electron microscopy
analysis, or (3) a segment was weighed and homogenized
in 10 volumes (w/v) of ice-cold buffer (300 mMsucrose,
1m
MEDTA, 10 mMNaN3,40mMTris-base and 40 mM
histidineatpH7.8)at0
◦Cina1mlglasshomogenizerwith
a glass pestle (Kontes Glass Industry, Vineland, NJ, USA).
Prior to homogenization the muscle sample was rinsed free
of contaminating blood by washing in ice-cold buffer. The
homogenate was divided into different portions and frozen
into liquid nitrogen for later analyses of Ca2+kinetics,
protein and myosin heavy chain (MHC) composition (see
below). In a few biopsies (out of 40 from both arm and leg
muscle) the sample portion was not enough to obtain all
three specimens.
SR vesicle Ca2+uptake and release measured
in crude muscle homogenate
The technique for measuring SR vesicle Ca2+uptake
and release was performed as previously described
in detail (Ørtenblad et al. 2000; Nielsen et al.
2007) (Fig. 1). Briefly, muscle homogenate (70 μl)
was mixed with 2 ml assay buffer (165 mMKC l,
22 mMHepes, 7.5 mMoxalate, 11 mMNaN3,5.5μM
N,N,N,N-tetrakis(2-pyridylmethyl)ethylenediamine
(TPEN), 20 μMCaCl2and 2 mMMgCl2(pH 7.0 at
37◦C)), and the reaction initiated by adding ATP to a
final concentration of 5 mM.[Ca
2+]wasdetermined
fluorometrically (20 Hz, Ratiomaster RCM, Photon
Technology International, Brunswick, NJ, USA) using the
fluorescent Ca2+indicator indo-1 (1 μM). When [Ca2+]
reached a plateau (nadir-Ca2+,i.e.[Ca
2+]attheendof
Ca2+uptake, Fig. 1 value c), SR Ca2+uptake was blocked
by adding cyclopiazonic acid (40 μM)andCa
2+release
was initiated by adding 4-chloro-m-cresol (5 mM), and
the fluorescence followed for at least 30 s.
Fluorescence was converted to free [Ca2+] (Ørtenblad
et al. 2000, Fig. 1 grey line) and raw data for [Ca2+]were
imported into Matlab version 7.0.1 (The MathWorks,
Natick, MA, USA) and mathematically analysed (Curve
Fitting Toolbox version 1.1.1; The MathWorks). Curve
fitting of Ca2+uptake was performed with data points
between a free [Ca2+] of 1000 nMand the free [Ca2+]
20 s prior to initiating Ca2+release (r2>0.99 for all data
sets, Fig. 1, dotted line between 0 and 330 s). The time
(τ) to reach 63% of the SR vesicle uptake (i.e. the initial
free [Ca2+] minus nadir-Ca2+) was calculated as 1/bfrom
the equation; y=ae−bt +c,whereyis the free [Ca2+],
tis time and a,band care constants assigned from
Matlab. There were no differences in the values of the
constants aand c(nadir-Ca2+)betweentrials,armand
leg, time or within same subject at various time points,
with aaveraging 1297 ±24 nMand nadir-Ca2+averaging
35 ±4n
M.
SR Ca2+release rate was obtained by mathematically
fitting the data points during the first 30 s of release to
the equation: y=x(1 – e−y(t−z)) (Fig. 1, dotted line from
330 s), back-extrapolate to nadir-Ca2+and the rate of
Ca2+release was determined as the derivative of the initial
release (Fig. 1, continuous line). The values obtained for
SR Ca2+-release rates are relative and therefore expressed
as arbitrary units of Ca2+min−1(g protein)−1). Due to the
inter-individual variation in SR Ca2+release rate, largely
owing to fibre-type differences (see Results), figures and
statistics are determined by relative SR Ca2+release and
uptake rates (%Pre). Assays of uptake and release rates of
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 715
Ca2+were performed in triplicate (a few in duplicate due
to limited tissue homogenate).
Protein content in the muscle homogenate was
measured in triplicate using a standard kit (Pierce BCA
protein reagent no. 23225). The homogenate protein
content averaged 11.1 ±0.2 mg ml−1, with no significant
difference between trials or time point, or in samples from
the same subject.
Fibre-type composition
Myosin heavy chain composition was analysed in the same
muscle homogenate as used for measurements of Ca2+
kinetics using gel electrophoresis as previously described
(Salviati et al. 1986) and modified for humans (Andersen &
Aagaard, 2000). Briefly, muscle homogenate (80 μl) was
mixed with 200 μl of sample buffer (10% glycerol, 5%
2-mercaptoethanol and 2.3% SDS, 62.5 mMTris and 0.2%
bromophenol blue at pH 6.8), boiled in a water bath at
100◦C for 3 min and loaded (10–40 μl) on a SDS-PAGE gel
(6% polyacrylamide (100:1, acrylamide:bis-acrylamide),
30% glycerol, 67.5 mMTris-base, 0.4% SDS, and 0.1 mM
glycine). Gels were run at 80 V for at least 42 h at 4◦Cand
MHC bands made visible by staining with Coomassie. The
gels were scanned (Linoscan 1400 scanner, Heidelberg,
Germany) and MHC bands densitometrically quantified
(Phoretix 1D, non-linear, Newcastle, UK). MHCII was
identified with Western blot using monoclonal antibody
(Sigma M 4276) with the protocol Xcell IITM (Invitrogen,
Carlsbad, CA, USA). Data are averages of three biopsies,
i.e.twofromoneleg/armandonefromtheotherleg/arm.
Biochemical determination of muscle glycogen
Muscles were frozen and later freeze-dried, dissected
free of non-muscle tissue, powdered and extracted with
perchloric acid as previously described (Harris et al. 1974).
Muscle glycogen was analysed from a separate portion of
the freeze-dried muscle according to Lowry & Passonneau
(1972).
Transmission electron microscopy (TEM) and
estimation of glycogen content and localization
A segment of the muscle was fixed for TEM analysis
as previously described in detail (Nielsen et al. 2010).
Briefly, the segment was fixed with 2.5% glutaraldehyde
in 0.1 Msodium cacodylate buffer (pH 7.3) for 24 h and
afterwards rinsed four times in 0.1 Msodium cacodylate
buffer. The segments were then post-fixed and stained
50 100 150 200 250 300 3500
200
400
600
800
1000
0
1200
τ= 1/
b
c
a
+
c
63.2% ↓
Rate of Ca2+ uptake fit;
nM Ca2+ = 1275 e-0.0326 + 36 (
r
2 = 0.99)
Rate of Ca2+ release fit;
nM Ca2+ . s-1 = 24.94
t
- 8179 (
r
2 = 0.97)
[Ca2+]free (nM)
Time (s)
Figure 1
Measurement of parameters associated with SR vesicle Ca2+uptake and Ca2+release from a representative [Ca2+]
trace (grey trace) obtained on a homogenate muscle sample from the arm after exercise using the assays described
in Methods.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
716 N. Ørtenblad and others J Physiol 589.3
with 1% osmium tetroxide (OsO4) and 1.5% potassium
ferrocyanide (K4Fe(CN)6)in0.1Msodium cacodylate
buffer for 90 min at 4◦C, the use of reduced osmium
tetroxide-containing potassium ferrocyanide favouring a
high electron density of glycogen particles (de Bruijn,
1973). After post-fixation the packed fibre segments were
rinsed twice in 0.1 Msodium cacodylate buffer at 4◦C,
dehydrated through graded series of alcohol at 4–20◦C,
infiltrated with graded mixtures of propylene oxide and
Epon at 20◦C, and embedded in 100% Epon at 30◦C.
Ultra-thin sections (∼60 nm) were cut using a Leica
Ultracut UCT ultramicrotome, and contrasted with uranyl
acetate and lead citrate. The sections were examined and
photographed in a pre-calibrated Philips EM 208 electron
microscope and a Megaview III FW camera.
Images were obtained at ×40,000 magnification in a
randomized systematic order to ensure unbiased results.
Three localizations of glycogen were defined: (1) the sub-
sarcolemmal space, (2) the intermyofibrillar space, and (3)
the intramyofibrillar space. The glycogen volume fraction
(VV) of each subpopulation was estimated as proposed
by Weibel (1980), where the effect of section thickness
was taken into account: VV=AA−t((1/π)BA−NA((t×
H)/(t+H))), where AAis the glycogen area fraction, tis
the section thickness (60 nm), BAis the glycogen boundary
length density, NAis the number of particles per area, and
His the average glycogen profile diameter.
Statistical analyses
All data are shown as means ±standard error of the
mean (S.E.M.). Data were analysed using unpaired
ttest, ANOVA when appropriate. Significant differences
between means were located using the Bonferroni post hoc
test. Correlation between two variables was tested with
Pearson’s correlation analysis using log transformed data.
Analyses were conducted in StatView 5.0 (SAS Institute,
Cary, NC, USA). The significance level was set toP≤0.05.
Results
Cross country race performance
The average race time for the ∼20 km race was
56 min 58 s ±1min 8s (range 52min 8s to 62min
30 s) for the ∼20 km race. Blood lactate measured
immediately after the race was 10.4 ±0.8 mmol l−1(range
7.7–14.1 mmol l−1).
Myosin heavy chain distribution
MHC distribution in m. vastus lateralis and m. triceps
brachii (distal part of the lateral head) revealed a clear over-
weight of MHCI in the leg muscle (58%), with the opposite
Table 1. Myosin heavy chain composition in arm and leg muscle
from elite cc-skiers
Fibre-type distribution (%)
MHCI MHCIIa MHCIIx
Leg (vastus lateralis) 58 ±241±21±0.4
Arm (triceps brachii) 40 ±3∗60 ±3∗0.4±0.2
Myosin heavy chain (MHC) composition from m. vastus lateralis
and m. triceps brachii (distal part of the lateral head). The
analysis was performed on three biopsies from each subject, i.e.
both left- and right-hand side muscles included, using SDS-PAGE.
∗Significantly different from leg muscle.
in the arm, revealing an average of 60% expression of
the MHCIIa isoform (Table 1). Two of the cc-skiers who
were very successful in sprint performance had more
than 70% of the arm fibres expressing MHCIIa, i.e. 71%
and 72%.
Glycogen content
Biochemically determined muscle glycogen was on average
reduced to 31 ±4% (from 540.2 to 167.1 mmol (kg
dry wt)−1in the arms and 71 ±10% (from 484.7 to
331.3 mmol (kg dry wt)−1) of the initial level in the legs
directly after the race (Fig. 2). Accordingly, there was a
pronounced higher muscle glycogen utilization in the
arms compared to the legs during the race (363 ±47 and
143 ±53 mmol (kg dry w t)−1,armsandlegs,respectively).
After 4 h recovery with CHO, arm glycogen was noticeably
recovered (59 ±5% of Pre) and fully recovered in the
leg muscle. In the absence of CHO during the first 4 h
recovery, muscle glycogen remained low (arm 29 ±2%
and leg 76 ±5%). However, after the remaining 18 h
recovery with CHO the muscle glycogen was restored to
pre-race levels in both the CHO and water trials (Fig. 2,
arm 85 ±7% and leg 101 ±3%).
SR vesicle function
There was a significant, on average 15 ±2%, decrease in
arm SR vesicle Ca2+releaseratefollowingtheexhaustivecc
race (from 4.7 ±0.4 to 3.9 ±0.4 μmol Ca2+(g protein)−1
min−1), with no significant difference between trials
(Fig. 3). However, there was no significant effect on leg
Ca2+release, but a significant difference between the
arm and leg release rates. In the arm, a 4 h recovery
with optimal CHO fully normalized the SR Ca2+release
rate. In the water trial, which omitted CHO during
the first 4 h of recovery, the muscle SR Ca2+release
rate remained reduced (on average, by 23 ±5%) and
significantly different from that of the 4 h recovery with
CHO (Fig. 3). In the water trial, SR Ca2+release rate
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 717
returned to normal values 22 h after the race, i.e. after
18 h with optimal CHO. There were no differences in leg
SR Ca2+release rates throughout the recovery period, in
both CHO and water trials.
In Pre biopsies there was a significant 34% higher SR
Ca2+release rate in arm compared to leg (4.7 ±0.4 vs.
3.5 ±0.3 μmol Ca2+(g protein)−1min−1, respectively).
When combining arm and leg data at rest, there was a
significant correlation between the MHCI distribution
and the SR vesicle Ca2+release rate (SR vesicle release
rate =6.913–0.054 %MHCI, r=0.34, P<0.01, n=19),
where SR vesicle release rate is given in μmol Ca2+
(g protein)−1min−1. This reveals that there is an
approximately 4.6 times higher SR Ca2+release rate
in MHCII fibres as compared to MHCI, i.e. release
rate of 6.9 and 1.5 with 100% MHCII and MHCI
fibres, respectively. This supports data from rodents
demonstrating an approximately 3-fold higher action
potential-induced peak rate of SR Ca2+release between the
fibre types (Baylor & Hollingworth, 2003). Accordingly,
with a 4.6 times higher release rate in MHCII fibres the
observed difference in the relative MHC composition of
the two muscles would be expected to give a 24% higher
Ca2+release rate in the arm, which is in line with present
data.
SR Ca2+uptake function was estimated as the
ATP-supported SR vesicle uptake and given as the τvalue,
calculated after curve fitting, defined as the time for the free
[Ca2+] to decrease by 63% of the initial free [Ca2+]. There
was no significant difference in the τvalue in arm following
the exhaustive race and neither in the 22 h recovery period
after the race. However, in leg muscle τwas significantly
increased by 23 ±12% immediately following the race
(i.e. SR Ca2+uptake decreased) from a resting value of
25.6 ±2.3s.SRCa
2+uptake in the leg muscles was fully
recovered within the first 4 h recovery with no difference
between trials.
Correlation between SR release rate and muscle
glycogen content
The study design enables the estimation of the possible
role of muscle glycogen in SR function since the muscle
milieu will be normalized after 4 h recovery, while muscle
Arm
Skeletal muscle glycogen content (mmol (kg dry wt)–1)
0
100
200
300
400
500
600
Leg
0
100
200
300
400
500
600
Skeletal muscle glycogen content (mmol (kg dry wt)–1)
*‡
4h waterCHO
Pre Post 4h 22h
4h waterCHO
*† ‡
*‡
*‡ #
*‡*‡*‡
Pre Post 4h 22h Pre Post 4h 22h Pre Post 4h 22h
Figure 2
Muscle glycogen was analysed biochemically in freeze-dried muscle from triceps brachii (arm) and vastus lateralis
(leg). Data are from before (Pre), immediately after (Post) and 4 h and 22 h after an approximately 1 h exhaustive
cross-country race. CHO denotes that the skiers received optimal CHO during the first 4 h, and water denotes that
for the first 4 h skiers only received water and no CHO. After the 4 h recovery, the skier assigned to the water trial
received the same CHO-enriched energy intake for the remaining time of the 22 h recovery, i.e. 18 h CHO for the
water trial. n=4–5.∗Significantly different from Pre; †significantly different from 4 h; ‡significantly different from
22 h; #significantly different from 4 h CHO trial.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
718 N. Ørtenblad and others J Physiol 589.3
glycogen levels vary in the recovery period, due to the
availability of CHO. In arm, which had a significant
decrease in the release rate and marked reduction in
muscle glycogen, there was a close association between
SR Ca2+release rate and muscle glycogen content (Figs 2
and 3). Thus, relatively low glycogen levels (Post and 4 h
water trial) are followed by an impaired SR Ca2+release
rate. In order to verify whether muscle glycogen levels are
associated with SR vesicle release rate at the individual
level, we correlated the individual data of SR release and
glycogen content from the cc-skiers. There was significant
logarithmic correlation (r2=0.29, n=67, P<0.0001)
between muscle total glycogen content and the relative
SR Ca2+release rate. When discriminating data from
arm and leg, only arm muscle (which exhibited very low
glycogen levels) revealed a significant correlation between
SR release rate and muscle glycogen (r2=0.41, n=34,
P<0.0001). Furthermore, a discrimination between time
points revealed a significant correlation after 4 h recovery
(r2=0.30, n=16, P=0.03, Fig. 4), and no significant
correlations at the other time points.
TEM-determined glycogen volume and association
with SR function
Since glycogen is heterogeneously distributed in the
muscle in distinct regions of the fibre (Marchand et al.
2002; Nielsen et al. 2010), we used TEM estimations of
glycogen volume in order to verify associations of the
discrete glycogen localization with the observed SR Ca2+
release rate. As previously stated, three localizations of
glycogen were defined in the fibres: (i) IMF, (ii) Intra, and
(iii) SS (Fig. 6) (Nielsen et al. 2010), with a relative volume
distribution of about 75–85%, 8–15% and 5–10% in the
intermyofibrillar, intramyofibrillar and subsarcolemmal
spaces, respectively (Marchand et al. 2007; Nielsen et al.
2010). The data showed a high correlation between
biochemically determined glycogen and TEM-estimated
glycogen (r2=0.57). When including all observations
(n=72) there was a significant correlation of SR Ca2+
release rate and all three subfractions of glycogen (IMF,
r2=0.15; P=0.001; Intra, r2=0.24; P<0.0001; SS,
r2=0.14, P=0.001). However, discrimination between
0
SR vesicle Ca2+ release rate (% control)
65
70
75
80
85
90
95
100
105
Pre Post 4h 22h
4h waterCHO
Pre Post 4h 22h Pre Post 4h 22h
4h waterCHO
Pre Post 4h 22h
0
SR vesicle Ca2+ release rate (% control)
65
70
75
80
85
90
95
100
105
Arm Leg
***#
Figure 3. SR vesicle Ca2+release rate in crude muscle homogenate
The SR vesicle Ca2+uptake and release were measured fluorometrically. Data were mathematically fitted using
mono-exponential equations before estimating the vesicle Ca2+release rate. Arm muscle SR Ca2+release rates
were on average decreased to 85 ±2% of initial values following the race. Four hours of recovery with CHO fully
normalized SR Ca2+release rate. However, in the absence of CHO during the first 4 h recovery the SR Ca2+release
rate remained low and reduced by 23 ±5%, returning to normal levels after the remaining 18 h recovery with
optimal CHO. There were no significant effects of exercise on the leg SR Ca2+release rate. n=4–5.∗Significantly
different from Pre; #significantly different from 22 h.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 719
r2 = 0.30
60
70
80
90
100
110
120
130
0 100 200 300 400 500
Glycogen (mmol (kg dry wt)–1)
SR Ca2+ release rate (% of Pre)
0
Figure 4. Correlation of SR Ca2+release rate and total
glycogen concentration in biopsies from cross-country skiers
obtained 4 h post exercise
Data points are included from the legs (circles) and arms (triangles)
from both CHO (filled symbols) and water (open symbols) trials. The
line indicates best fit of all the data points (r2= 0.30, P= 0.03,
n= 16).
time points revealed a significant correlation between
Intra glycogen and SR Ca2+release rate 4 h post
(r2=0.23; P=0.04, Fig. 5B) and there were no significant
correlations at the other time points or subfractions. If
the correlation analysis is further stratified by limbs no
correlations were present using this small sample size
(n=9–10). Thus a small fraction (10–15%) of glycogen
that is localized inside the myofibrils, near the SR-voltage
sensors at the triad in the I-band, appears to be most
closely associated with the SR Ca2+release rate.
Discussion
In the present study, we have investigated the role of intense
1 h exercise and glycogen availability on the SR function
in the arm and leg muscles of highly trained cc-skiers.
The main findings were that following the exercise, the
SR Ca2+release rate was only significantly decreased in
the arms and that this coincided with lower glycogen
content when compared to the legs. The role of glycogen
availability was strengthened by the finding that the
diminished SR vesicle Ca2+release rate was not improved
if glycogen was maintained low in the recovery period
by the absence of CHO intake indicating a direct role
for glycogen in regulating SR Ca2+release rate. Further,
regression analysis of subcellular localizations of glycogen
and the SR Ca2+release rate suggested that only glycogen
localized inside the myofibrils (intramyofibrillar glycogen)
was associated with SR Ca2+release rate. The results
raise the possibility that intramyofibrillar glycogen is a
prerequisite for normal SR Ca2+release, and that feedback
regulation from glycogen storage deposits located inside
the myofibrils may exist. Thus, glycogen may regulate the
cytosolic levels of Ca2+and, in turn, the energy utilization
of skeletal muscle.
SR function in fatigue
It is clear that reduced SR function in general, and SR
Ca2+release in particular, is an important contributor to
muscle fatigue during a variety of exercise types (Favero,
1999; Allen et al. 2008). The demonstration of intrinsic
dysfunctions of the SR as an important mechanism of
fatigue is gained from both studies at the single cell level
(Chin & Allen, 1997; Allen et al. 2008), isolated muscles
(Ward et al. 1998; Ørtenblad et al. 2000), and whole
body exercise (Byrd et al. 1989; Gollnick et al. 1991;
Duhamel et al. 2006a,b). Thus, the present findings in
highly trained athletes are consistent with previous studies,
r2 = 0.08
50
60
70
80
90
100
110
120
130
0 0.01 0.02 0.03 0.04
IMF Glycogen (μm3 μm-3)
0 0.002 0.004 0.006
Intra Glycogen (μm3 μm-3)
0.00 0.02 0.04 0.06 0.08 0.10
SS Glycogen (μm3 μm-2)
SR Ca2+ release rate (% of Pre)
A
0
r2 = 0.23 r2 = 0.12
50
60
70
80
90
100
110
120
130
SR Ca2+ release rate (% of Pre)
B
0 50
60
70
80
90
100
110
120
130
SR Ca2+ release rate (% of Pre)
C
0
Figure 5. Correlation of TEM-determined subfractions of glycogen and SR Ca2+release rate
A, IMF glycogen; B, Intra glycogen; C, SS glycogen. Data points are included from the legs (circles) and arms
(triangles) from both CHO (filled symbols) and water (open symbols) trials. Lines indicate best fits of all the data
points (A:r2= 0.08, P= 0.23, n= 19; B:r2= 0.23, P= 0.04, n= 19; C,r2= 0.12, P= 0.14, n= 19).
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
720 N. Ørtenblad and others J Physiol 589.3
showing an exercise-induced impairment in the muscle
Ca2+regulation following 1 h of exhaustive exercise. Direct
measurements of SR Ca2+uptake and release rates can
only be obtained in SR vesicle preparations and these
studies have demonstrated exercise-related decreases in
both or either of these properties. Several studies have
reported an up to 40% decrease in the SR vesicle Ca2+
release rate following both intense and prolonged exercise
in humans (Hill et al. 2001; Li et al. 2002; Leppik
et al. 2004; Duhamel et al. 2006a). Currently, several
mechanisms are likely candidates for explaining the in
vivo reduction in SR function, depending on fatigue mode
and intracellular environment, e.g. low ATP, elevated
Mg2+,Ca
2+, intracellular phosphate, or reactive oxygen
species. However, measurements of SR vesicle function are
carried out in vitro under apparently optimal conditions,
and demonstrate that once the SR is removed from
the intracellular environment and not under normal
voltage sensor control, changes in function persist. This,
however, does not imply that the depressed SR vesicle
function unequivocally is linked to a decreased rate of
Ca2+release during normal in vivo excitation–contraction
coupling. One factor that appears to affect SR vesicle
function is the glycogen content of the skeletal muscle
cell. In turn, Duhamel and colleagues (Duhamel et al.
2006a,c), using the same method as in the present
study, have shown that there is a clear association
between muscle glycogen content and SR vesicle function
during prolonged exercise, when starting exercise with
high and low glycogen levels, respectively. Furthermore,
present data on human muscles obtained immediately
after exercise and during the recovery period following
exercise, support the data from both single fibres and
muscle bundles that demonstrate that muscle force and
tetanic [Ca2+]iare associated with muscle glycogen
content (Chin et al. 1997; Kabbara et al. 2000; Helander
et al. 2002) in the recovery phase after fatiguing contrac-
tions. Together, these data suggest that in elite trained
humans the change in SR function is also associated with
fatigue and that impaired SR function has a component
that is glycogen dependent. There was no significant
decrease in the SR vesicle Ca2+release rate in leg muscle
following exhaustive exercise and there was a significantly
greater decrease in the arm vesicle release rate compared
to that of the leg (P<0.05). This may be unexpected
and explained by a lack of power, either due to a limited
number of subjects and SR data (n=9) or the different
fibre-type distribution in the arm muscles compared to
leg muscles (Table 1). Thus, assuming that the MHCII
fibres are more prone to impaired SR Ca2+release, the
arm muscles would be more affected than leg muscles, as
well as if the MHCII fibres were more active in the arms
than in the legs.
In the present study of highly trained cc-skiers, the SR
Ca2+uptake rate in leg muscle was significantly reduced
after exercise, while fully recovered after a 4 h recovery.
However, we did not observe an effect on SR vesicle Ca2+
uptake rate following exercise in the arm. When viewed
as a whole, most studies have reported a reduced SR
vesicle Ca2+uptake following exercise (Luckin et al. 1991;
Duhamel et al. 2006a), though SR Ca2+uptake has also
been reported to be unaltered (Dossett-Mercer et al. 1994,
1995). These discrepancies may be related to the exercise
mode and training status. In term of glycogen, our results
suggest that in elite cc-skiers whole-muscle glycogen is not
associated with the SR Ca2+uptake rate, whereas it could
be speculated that the exercise mode of the leg muscle may
determine the susceptibility of the SR Ca2+pump to the
exercise.
Association between SR and the glycogenolytic
complex
An important assumption for the present, and previous,
conclusions is that the SR vesicle used for the in vitro
measurements of SR function is associated with glycogen.
Therefore, even though SR dysfunction is evident when
it is removed from the intracellular milieu and evaluated
in constant and optimal conditions, it is important to
consider the physiological relevance of glycogen in the SR
vesicle preparation. Previous studies have demonstrated
a strong binding between SR membrane and glycogen
and glycogen-regulating proteins, and the existence of
a glycogenolytic complex consisting of glycogen and
enzymes involved in its metabolism is nowwell established
(Wanson & Drochmans, 1972; Entman et al. 1980;
Kruszynska et al. 2001; Shearer & Graham, 2004). This
association of glycogen, related enzymes and regulating
proteins substantiates the existence of a functionally
independent glycogenolytic complex, which is involved in
the control of its own glycolytic metabolism. Furthermore,
it has been demonstrated that part of this glycogenolytic
complex can be physically associated with the SR (Wanson
& Drochmans, 1972; Entman et al. 1980; Cuenda et al.
1995; Xu & Becker, 1998; Lees et al. 2001, 2004).
Thus, Entman and collegues have demonstrated that
the SR–glycogenolytic complex is a highly specific,
functionally defined compartment for phosphorylase
regulation and that this is a compartmented system for
phosphorylase activation controlled by SR calcium flux
(Entman et al. 1980). Importantly, the association between
SR and the glycogenolytic complex is apparently dynamic,
changing with the metabolic state of the muscle cell (Xu
& Becker, 1998; Kruszynska et al. 2001; Lees et al. 2001).
Taken together, these observations provide evidence for
an integrated and dynamic association between the SR
and part of the glycogenolytic complex, which appears
to be essential for SR function and persists in SR vesicle
preparations.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 721
Association of glycogen and SR Ca2+release rate
The present data demonstrate a 15% decrease in
the SR Ca2+release rate in arm after exhaustive
exercise, associated with low glycogen levels (<200 mmol
(kg dry wt)−1). Further, when muscle glycogen was kept
low by omitting CHO in the recovery period, the SR
Ca2+release rate remained impaired. Importantly, the SR
Ca2+release rate recovered and was significantly different
from the water trial when the athletes received optimal
CHO during the 4 h recovery period and muscle glycogen
was increased to 283 ±8 mmol (kg dry wt)−1(59 ±5%
of Pre). These measurements after 4 h recovery eliminate
parallel changes in a variety of candidates that traditionally
explain the reduction in SR function when fatigued, and
strongly indicate that low muscle glycogen per se affects
SR Ca2+release rate.
A critical level of muscle glycogen below which SR
function is impaired has been suggested (Duhamel et al.
2006c). This concept arises from the observations that:
(1) a small decrease in muscle glycogen does not impair
muscle Ca2+regulation (Chin & Allen, 1997; Duhamel
et al. 2006c), (2) under conditions with low starting muscle
glycogen the impaired Ca2+regulation is accelerated
(Chin & Allen, 1997; Helander et al. 2002), and (3)
exercise-induced reductions in SR function are observed
much later during exercise when starting with high
glycogen compared to low glycogen during whole body
exercise (Duhamel et al. 2006c). Thus it was observed that
the SR vesicle Ca2+release rate was significantly depressed
at muscle glycogen levels below an apparent critical
level of 300 mmol (kg dry wt)−1, which was attained
after ∼30 min and ∼60 min of exercise, with low and
high starting glycogen levels, respectively (Duhamel et al.
2006c). These data are very much in line with the pre-
sent observation of a decrease in SR Ca2+release rate
associated with very low glycogen levels (<200 mmol
(kg dry wt)−1). Importantly, when muscle glycogen was
kept low by omitting CHO in the recovery period,
SR Ca2+release rate remained impaired, while the SR
Ca2+release rate recovered and was significantly different
from that in the water trial, when muscle glycogen was
283 ±8 mmol (kg dry wt)−1.Fromthepresentdataand
previously published data (Duhamel et al. 2006c)we
suggest a critical level of muscle glycogen of 280–300 mmol
(kg dry wt)−1,belowwhichtheSRCa
2+release rate is
impaired. The idea of a critical level of total glycogen
can, however, be challenged by the present results using
TEM, showing that only intramyofibrillar glycogen is
associated with SR Ca2+release rate. Nevertheless, the
logarithmic correlation between Intra glycogen and SR
Ca2+release rate 4 h post exercise indicates that a critical
level for the Intra glycogen is necessary for optimal SR Ca2+
release.
Glycogen localization and the role of distinct pools
Both qualitative and quantitative studies have shown that
repeated contractions mediate an uneven breakdown of
glycogen fractions, raising the likely possibility that the
various glycogen pools, or even granula, may act as
independent metabolic units with specialized functions,
resulting in regulatory or metabolic compartments with
distinct characteristics and functions (Sj¨
ostr¨
om et al.
1982a;Fridenet al. 1985, 1989; Marchand et al.
2007; Nielsen et al. 2009). In line with this, we have
recently shown that in mechanically skinned fibres,
the distinct pool of intramyofibrillar glycogen was
positively correlated with the fatigue resistance capacity.
These results demonstrate that the distinct subcellular
populations of glycogen have different roles in contracting
single muscle fibres and that muscle fatigueability is partly
controlled by the availability of intramyofibrillar glycogen
(Nielsen et al. 2009). Further, the intramyofibrillar pool is
recognized as being preferentially restored after prolonged
exercise (Marchand et al. 2007). Here we demonstrate for
the first time, in whole muscles from elite cc-skiers, that it
is the particular subfraction of intramyofibrillar glycogen
which is significantly associated with the SR Ca2+release
rate (r2=0.23, Fig. 5). The fact that intramyofibrillar
glycogen is not physically associated with SR suggests that
the link between low glycogen and SR Ca2+release rate
is mediated through a signalling cascade. Importantly,
in this study, using trained athletes and prolonged
exercise, the biochemically determined glycogen (whole
muscle glycogen) and TEM-estimated glycogen were
relatively closely correlated. This relationship explains the
observed association between biochemically determined
glycogen and the SR Ca2+release rate despite the
apparent significance of the particular subfraction of intra-
myofibrillar glycogen on the SR Ca2+release rate. The
finding of a higher correlation coefficient when the SR
Ca2+release rate is related to biochemically determined
glycogen (r2=0.30) rather than to TEM-estimated
Intra glycogen (r2=0.23) may suggest that biochemical
glycogen is the best predictor of SR Ca2+release rate.
However, the higher r2value is due to a lower sample size
in biochemically determined glycogen and a correction for
the sample size reveals similar r-values (data not shown)
by the two methods.
Coupling mechanisms between intramyofibrillar
glycogen and SR Ca2+release
The vast majority of intramyofibrillar glycogen is
specifically located in the I-band of the sarcomere,
which is close to the triads, with the voltage sensor
and SR Ca2+release channel (RyR). In line with the
idea of a compartmentalized energy transfer between
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
722 N. Ørtenblad and others J Physiol 589.3
glycolytic-derived ATP and energy-controlled processes,
the role of intramyofibrillar glycogen could be to
provide glycolytic intermediates to the triads maintaining
an optimal coupling between voltage sensors and SR
RyRs. Such an apparent coupling has previously been
demonstrated between both glycolysis and the skeletal
muscle transverse-tubule Na+,K+-pumps (Dutka & Lamb,
2007) and with the existence of a sequential glycolytic
reaction compartmentalized in the triads, where the
glycolytic-derived ATP formation appears not to be in
equilibrium with the bulk ATP (Han et al. 1992; Less et al.
2004). The triad is located in a very restricted space formed
by the terminal cisternae of the SR, which lie alongside
the T-system, and it is thereby clearly separated from
mitochondria and intermyofibrillar glycogen. Thus, a high
and constant myoplasmic energy status does not imply that
the energy status is constant in the very restricted space in
the triadic gap (12 nm and limited by the SR) with a high
local energy turnover during repetitive contractions. The
triadic gap is physically remote from mitochondria, lipids,
glycogen and other constituents of the intermyofibrillar
space (Fig. 6B). Thus, access of glycolytic intermediates,
from the breakdown of glycogen to the triad, could be
speculated to be favoured from intramyofibrillar glycogen
localized in the I-band near the triads, but not necessarily
physically associated with the SR. On the contrary, inter-
myofibrillar glycogen does not seem to be directly located
next to the triad and both intermyofibrillar and sub-
sarcolemmal glycogen might have limited access to the
triad gap, which is covered by the SR. In line with this,
several glycolytic enzymes are present in the triads, and do
not seem to be in direct equilibrium with the bulk enzymes
(Han et al. 1992). This association between the SR, the
enzymes and the connection to intramyofibrillar glycogen
apparently persists with the SR vesicle preparation as used
in the present study. The present demonstration of a
correlation between intramyofibrillar glycogen and the SR
Ca2+release rate suggests that compartmentalized energy
transfer through glycolysis or signalling via glycolytic
intermediates is essential for normal muscle function
and may explain the long time recognized importance of
muscle glycogen on performance, during both prolonged
and high intensity intermittent exercise.
Conclusions
This study demonstrates, in human elite cc-skiers, a close
association between low glycogen (less than about 50%
of control) and impaired SR Ca2+release. This strongly
indicates that glycogen content modulates the SR Ca2+
release rate, which is in agreement with the emerging
concept of a structural role for glycogen, meaning it
has a much more diverse functional role than just an
energy reserve. Moreover, transmission electron micro-
scopy analysis revealed that the distinct fraction of
glycogen localized in the intramyofibrillar space, was
highly significantly correlated with the SR Ca2+release
rate. Thus, the depletion of glycogen during prolonged,
exhausting exercise may contribute to fatigue by causing
a decreased SR Ca2+release rate. Further, the pre-
sent findings suggest that the link between glycogen
and SR Ca2+release rate is compartmentalized to
Figure 6. TEM images showing the subcellular localization of skeletal muscle glycogen
Both images are from a pre-exercise biopsy of arm m. triceps brachii.A,overview of part of fibre showing the
myofibrillar (Myo) space and subsarcolemmal (SS) space. At the top of the image an adjacent fibre with a nucleus
(N) can be seen and in the interstitial space between the two fibres a capillary (C) is found. All the grey structures
in the fibre are mitochondria and black areas represent spaces filled with glycogen. The black box indicates the
position of the image at higher magnification shown in B.Scalebar=5μm. Original ×5000 magnification. B,
typical localization pattern of IMF and Intra glycogen in skeletal muscle. Glycogen particles are seen as black dots.
Z, Z-line; M, M-band. Scale bar = 0.5 μm. Original ×40,000 magnification.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 723
intramyofibrillar glycogen particles and may provide
further insight to explain the long recognized importance
of muscle glycogen on performance during both
prolonged and high intensity intermittent exercise.
Perspectives
The importance of carbohydrate as a fuel during
exercise has been recognized since the beginning of last
century. Using the muscle biopsy technique, it was later
demonstrated that the importance of carbohydrate can
be explained by a close association between the muscle
glycogen concentration and time to exhaustion. We have
now demonstrated that low muscle glycogen is associated
with an impairment of SR Ca2+release, leading to
fatigue. Thus, the depletion of glycogen during prolonged,
exhausting exercise may contribute to fatigue by causing
decreased SR Ca2+release. We show here that the specific
localization of glycogen in the intramyofibrillar space is
important for a normal SR Ca2+release rate. These data
provide strong indications for the importance of the sub-
cellular arrangement of glycogen in muscle fibres and the
possible existence of a compartmentalized energy trans-
fer or signal transduction from intramyofibrillar glycogen
particles to the restricted area of the triad junction that
modulates the SR Ca2+release rate.
References
Allen DG, Lamb GD & Westerblad H (2008). Skeletal muscle
fatigue: cellular mechanisms. Physiol Rev 88, 287–332.
Andersen JL & Aagaard P (2000). Myosin heavy chain IIX
overshoot in human skeletal muscle. Muscle Nerve 23,
1095–1104.
Barnes M, Gibson LM & Stephenson DG (2001). Increased
muscle glycogen content is associated with increased
capacity to respond to T-system depolarisation in
mechanically skinned skeletal muscle fibres from the rat.
Pflugers Arch 442, 101–106.
Baylor SM & Hollingworth S (2003). Sarcoplasmic reticulum
calcium release compared in slow-twitch and fast-twitch
fibres of mouse muscle. JPhysiol 551, 125–138.
Bergstr¨
om J, Hermansen L, Hultman E & Saltin B (1967). Diet,
muscle glycogen and physical performance. Acta Physiol
Scand 71, 140–150.
Byrd SK, Bode AK & Klug GA (1989). Effects of exercise of
varying duration on sarcoplasmic reticulum function. JAppl
Physiol 66, 1383–1389.
Calbet JA, Holmberg HC, Rosdahl H, van Hall G,
Jensen-Urstad M & Saltin B (2005). Why do arms extract less
oxygen than legs during exercise? Am J Physiol Regul Integr
Comp Physiol 289, R1448–R1458.
Chin ER & Allen DG (1997). Effects of reduced muscle
glycogen concentration on force, Ca2+release and
contractile protein function in intact mouse skeletal muscle.
JPhysiol498, 17–29.
Chin ER, Balnave CD & Allen DG (1997). Role of intracellular
calcium and metabolites in low-frequency fatigue of mouse
skeletal muscle. Am J Physiol Cell Physiol 272, C550–C559.
Christensen EH & Hansen O (1939). Arbeitsfahigkeit und
ernahrung. Skand Arch Physiol 81, 160–171.
Cuenda A, Nogues M, Henao F & Gutierrez-Merino C (1995).
Interaction between glycogen phosphorylase and
sarcoplasmic reticulum membranes and its functional
implications. J Biol Chem 270, 11998–12004.
de Bruijn WC (1973). Glycogen, its chemistry and morphologic
appearance in the electron microscope. I. A modified OsO4
fixative which selectively contrasts glycogen. J Ultrastruct Res
42, 29–50.
Dossett-Mercer J, Green H, Chin ER & Grange F (1995).
Failure of short term stimulation to reduce sarcoplasmic
reticulum Ca2+-ATPase function in homogenates of rat
gastrocnemius. MolCellBiochem146, 23–33.
Dossett-Mercer J, Green HJ, Chin E & Grange F (1994).
Preservation of sarcoplasmic reticulum Ca2+-sequestering
function in homogenates of different type composition
following sprint activity. Can J Physiol Pharmacol 72,
1231–1237.
Duhamel TA, Green HJ, Perco JG & Ouyang J (2006a).
Comparative effects of a low-carbohydrate diet and exercise
plus a low-carbohydrate diet on muscle sarcoplasmic
reticulum responses in males. Am J Physiol Cell Physiol 291,
C607–C617.
Duhamel TA, Green HJ, Perco JG & Ouyang J (2006b). Effects
of prior exercise and a low-carbohydrate diet on muscle
sarcoplasmic reticulum function during cycling in women.
JApplPhysiol 101, 695–706.
Duhamel TA, Perco JG & Green HJ (2006c). Manipulation of
dietary carbohydrates after prolonged effort modifies muscle
sarcoplasmic reticulum responses in exercising males. Am J
Physiol Regul Integr Comp Physiol 291, R1100–R1110.
Dutka TL & Lamb GD (2007). Na+-K+pumps in the
transverse tubular system of skeletal muscle fibers
preferentially use ATP from glycolysis. Am J Physiol Cell
Physiol 293, C967–C977.
Entman ML, Keslensky SS, Chu A & Van Winkle WB (1980).
The sarcoplasmic reticulum-glycogenolytic complex in
mammalian fast twitch skeletal muscle. Proposed in vitro
counterpart of the contraction-activated glycogenolytic pool.
J Biol Chem 255, 6245–6252.
Favero TG (1999). Sarcoplasmic reticulum Ca2+release and
muscle fatigue. JApplPhysiol 87, 471–483.
Favero TG, Pessah IN & Klug GA (1993). Prolonged exercise
reduces Ca2+release in rat skeletal muscle sarcoplasmic
reticulum. Pflugers Arch 422, 472–475.
Friden J, Seger J & Ekblom B (1985). Implementation of
periodic acid-thiosemicarbazide-silver proteinate staining
for ultrastructural assessment of muscle glycogen utilization
during exercise. Cell Tissue Res 242, 229–232.
Friden J, Seger J & Ekblom B (1989). Topographical localization
of muscle glycogen: an ultrahistochemical study in the
human vastus lateralis. Acta Physiol Scand 135, 381–391.
Gollnick PD, Korge P, Karpakka J & Saltin B (1991).
Elongation of skeletal muscle relaxation during exercise is
linked to reduced calcium uptake by the sarcoplasmic
reticulum in man. Acta Physiol Scand 142, 135–136.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
724 N. Ørtenblad and others J Physiol 589.3
Green HJ (1991). How important is endogenous muscle
glycogen to fatigue in prolonged exercise? Can J Physiol
Pharmacol 69, 290–297.
Han JW, Thieleczek R, Varsanyi M & Heilmeyer LM Jr (1992).
Compartmentalized ATP synthesis in skeletal muscle triads.
Biochemistry 31, 377–384.
Harris RC, Hultman E & Nordesj¨
o LO (1974). Glycogen,
glycolytic intermediates and high-energy phosphates
determined in biopsy samples of musculus quadriceps
femoris of man at rest. Methods and variance of values.
Scand J Clin Lab Invest 33, 109–120.
Helander I, Westerblad H & Katz A (2002). Effects of glucose
on contractile function, [Ca2+]i, and glycogen in isolated
mouse skeletal muscle. Am J Physiol Cell Physiol 282,
C1306–C1312.
Hill CA, Thompson MW, Ruell PA, Thom JM & White MJ
(2001). Sarcoplasmic reticulum function and muscle
contractile character following fatiguing exercise in humans.
JPhysiol531, 871–878.
Holmberg HC, Lindinger S, St¨
oggl T, Eitzlmair E &
M¨
uller E (2005). Biomechanical analysis of double
poling in elite cross-country skiers. Med Sci Sports Exerc 37,
807–818.
Kabbara AA, Nguyen LT, Stephenson GM & Allen DG (2000).
Intracellular calcium during fatigue of cane toad skeletal
muscle in the absence of glucose. JMuscleResCellMotil21,
481–489.
Krogh A & Lindhard J (1920). The relative value of fat and
carbohydrate as sources of muscular energy: with appendices
on the correlation between standard metabolism and the
respiratory quotient during rest and work. Biochem J 14,
290–363.
Kruszynska YT, Ciaraldi TP & Henry RR (2001). In Handbook
of Physiology. A Critical, Comprehensive Presentation of
Physiological Knowledge and Concepts,section7,The
Endocrine System, ed. Jefferson LS & Cherrington A, pp.
579–607. Oxford University Press, New York.
Lees SJ, Franks PD, Spangenburg EE & Williams JH (2001).
Glycogen and glycogen phosphorylase associated with
sarcoplasmic reticulum: effects of fatiguing activity. JAppl
Physiol 91, 1638–1644.
Lees SJ & Williams JH (2004). Skeletal muscle sarcoplasmic
reticulum glycogen status influences Ca2+uptake supported
by endogenously synthesized ATP. Am J Physiol Cell Physiol
286, C97–C104.
Leppik JA, Aughey RJ, Medved I, Fairweather I, Carey MF &
McKenna MJ (2004). Prolonged exercise to fatigue in
humans impairs skeletal muscle Na+-K+-ATPase activity,
sarcoplasmic reticulum Ca2+release, and Ca2+uptake.
JApplPhysiol 97, 1414–1423.
Li JL, Wang XN, Fraser SF, Carey MF, Wrigley TV & McKenna
MJ (2002). Effects of fatigue and training on sarcoplasmic
reticulum Ca2+regulation in human skeletal muscle. JAppl
Physiol 92, 912–922.
Lowry OH & Passonneau JV (1972). AFlexibleSystemof
Enzymatic Analysis. Academic Press, New York.
Luckin KA, Favero TG & Klug GA (1991). Prolonged exercise
induces structural changes in SR Ca2+-ATPase of rat muscle.
Biochem Med Metab Biol 46, 391–405.
Marchand I, Chorneyko K, Tarnopolsky M, Hamilton S,
Shearer J, Potvin J & Graham TE (2002). Quantification
ofsubcellularglycogeninrestinghumanmuscle:granule
size, number, and location. JApplPhysiol93,
1598–1607.
Marchand I, Tarnopolsky M, Adamo KB, Bourgeois JM,
Chorneyko K & Graham TE (2007). Quantitative assessment
ofhumanmuscleglycogengranulessizeandnumberin
subcellular locations during recovery from prolonged
exercise. JPhysiol580, 617–628.
Nielsen J, Mogensen M, Vind BF, Sahlin K, Højlund K,
Schrøder HD & Ørtenblad N (2010). Increased
subsarcolemmal lipids in type 2 diabetes: effect of training
on localization of lipids, mitochondria, and glycogen in
sedentary human skeletal muscle. Am J Physiol Endocrinol
Metab 298, E706–E713.
Nielsen J, Schrøder HD, Rix CG & Ørtenblad N (2009).
Distinct effects of subcellular glycogen localization on tetanic
relaxation time and endurance in mechanically skinned rat
skeletal muscle fibres. JPhysiol587, 3679–3690.
Nielsen JS, Sahlin K & Ørtenblad N (2007). Reduced
sarcoplasmic reticulum content of releasable Ca2+in rat
soleus muscle fibres after eccentric contractions. Acta Physiol
(Oxf) 191, 217–228.
Ørtenblad N, Sjogaard G & Madsen K (2000). Impaired
sarcoplasmic reticulum Ca2+release rate after fatiguing
stimulation in rat skeletal muscle. JApplPhysiol89,
210–217.
Salviati G, Betto R, Danieli-Betto D, Biasia E, Serena M, Mini
M & Scarlato G (1986). Myosin light chains and muscle
pathology. Neurology 36, 693–697.
Shearer J & Graham TE (2004). Novel aspects of skeletal muscle
glycogen and its regulation during rest and exercise. Exerc
Sport Sci Rev 32, 120–126.
Sj¨
ostr¨
om M, Friden J & Ekblom B (1982a). Fine structural
details of human muscle fibres after fibre type specific
glycogen depletion. Histochemistry 76, 425–438.
Sj¨
ostr¨
om M, Neglen P, Friden J & Eklof B (1982b). Human
skeletal muscle metabolism and morphology after
temporary incomplete ischaemia. Eur J Clin Invest 12,
69–79.
Stephenson DG, Nguyen LT & Stephenson GM (1999).
Glycogen content and excitation–contraction coupling in
mechanically skinned muscle fibres of the cane toad.
JPhysiol519, 177–187.
Wanson JC & Drochmans P (1972). Role of the sarcoplasmic
reticulum in glycogen metabolism. Binding of
phosphorylase, phosphorylase kinase, and primer complexes
to the sarcovesicles of rabbit skeletal muscle. J Cell Biol 54,
206–224.
Ward CW, Spangenburg EE, Diss LM & Williams JH (1998).
Effects of varied fatigue protocols on sarcoplasmic reticulum
calcium uptake and release rates. Am J Physiol Regul Integr
Comp Physiol 275, R99–R104.
Weibel ER (1980). Stereological Methods,vol.2,Theoretical
Foundations . Academic Press, London.
Xu KY & Becker LC (1998). Ultrastructural localization of
glycolytic enzymes on sarcoplasmic reticulum vesticles.
J Histochem Cytochem 46, 419–427.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
J Physiol 589.3 Muscle glycogen content and SR function in elite cross-country skiers 725
Author contributions
The experiments were performed at the Institute of Sports
Science and Clinical Biomechanics (muscle analysis) and
Institute of Pathology, Faculty of Health Science (electron
microscopy pictures), University of Southern Denmark,
DK-5230 M, Denmark, and the Swedish Winter Sports
Research Centre, Department of Health Sciences, Mid Sweden
University, ¨
Ostersund, Sweden (exercise and testing). All authors
contributed to the conception and design of the experiments,
collection, analysis and interpretation of data, and drafting the
article or revising it critically for important intellectual content.
All authors approved the final version of the manuscript.
Acknowledgements
The study was supported by The Danish Ministry of Culture,
the Committee on Sports Research (TKIF2005-049) and TEAM
Denmark (06-40215/3). The authors have no conflicts of interest
to disclose.
C
2011 The Authors. Journal compilation C
2011 The Physiological Society
