Comparison of traditional measurements with macroglycogen
and proglycogen analysis of muscle glycogen
K. B. ADAMO AND T. E. GRAHAM
Human Biology and Nutritional Sciences, University of Guelph,
Guelph, Ontario, Canada N1G 2W1
metabolic pools; glycogenin; measurement techniques; carbohydrate; biopsy
has been used for many years to
measure muscle metabolites, including glycogen, and it
is vital that the methods employed give an accurate
representation of the muscle glycogen stores. Glycogen
is found in granules, and thus there is the potential for
a high degree of variability among biopsy samples. An
electron-microscopic study of glycogen granules by
Friden et al. (5) demonstrated that there are two
different particle-sized populations in human skeletal
muscle. These two separate populations of glycogen
particles were stored in five different locations within
the muscle cell, and the authors speculated that these
populations could be two forms of the glycogen molecule
that were utilized at different times from distinct
locations. Despite these potential variables, Essen and
Henriksson (4), Harris et al. (6), and Hultman (9) have
shown that, in human muscle, the procedure is very
reliable and that muscle biopsy samples are representative of the whole muscle.
Traditionally, there have been two methods for measuring glycogen, either by acid hydrolysis (AC) (15) or
by enzymatic hydrolysis (EZ) (3). Although it has been
stated that the variability of the analytic technique is
about 610–13 mmol glucosyl units/kg dry weight (dw)
THE BIOPSY TECHNIQUE
908
(6), no detailed comparison of AC vs. EZ has been
published determining whether the methods are comparable. Bergmeyer (3) has suggested that EZ analysis of
purified glycogen gives values that are 5% higher than
those obtained by using AC, whereas Passonneau and
Lauderdale (15) claimed that the two methods give
equivalent results. None of above-mentioned authors
provided data to substantiate his claim. In the only
direct comparison of AC and EZ that we are aware of,
Jansson (10) performed the AC and EZ methods in
human muscle over a wide concentration range and
stated that there was no systematic difference between
the values obtained by using either method. However,
no data were provided to validate this finding.
Before Jansson’s study (10) it was common for glycogen measurements to be made in the precipitate that
had been pretreated with perchloric acid (PCA) (11, 15,
16). This method was thought to make the most efficient use of a muscle sample by allowing for the
precipitation of proteins and extraction of PCA-soluble
metabolites while also permitting the determination of
glycogen in the same piece of tissue. However, it has
been reported that only a portion of the total glycogen
was extracted during PCA treatment (7, 9, 11, 17).
Jansson (10) compared this method to AC and EZ and
documented that ,15–25% of the glycogen was PCA
soluble and hence any measurement of glycogen only
from the precipitate would underestimate the total.
Jansson’s findings discouraged the analysis of PCA
precipitate to determine total glycogen but did not
stimulate further research regarding the possibility
that these two fractions represented two physiological
pools.
Recently, investigators (1, 12, 14), while identifying
and studying glycogenin, the protein core of the glycogen molecule, realized that there were two pools of
glycogen in rodent skeletal muscle and other tissues
such as liver and heart. This work used only rodent
resting muscle and had not investigated exercise metabolism or human tissue.
The above-mentioned studies determined that one of
the fractions was of smaller molecular weight (400,000)
and was relatively rich in protein, prompting use of the
name proglycogen (PG). The other type, a larger molecule, was what is recognized as ‘‘classic’’ glycogen or
macroglycogen (MG; mol wt 10,000,000). The two forms
differ in the ratio of protein to carbohydrate. PG was
found to precipitate in trichloracetic acid because of its
10% protein component (12). MG, with a protein content of only 0.35%, was soluble.
No one has systematically compared the two traditional methods, nor have these methods ever been
compared with the MG/PG determination for glycogen.
0161-7567/98 $5.00 Copyright r 1998 the American Physiological Society
http://www.jap.org
Downloaded from http://jap.physiology.org/ by 10.220.33.5 on June 18, 2017
Adamo, K. B., and T. E. Graham. Comparison of traditional measurments with macroglycogen and proglycogen
analysis of muscle glycogen. J. Appl. Physiol. 84(3): 908–913,
1998.—Traditionally, there have been two methods for measuring total muscle glycogen (Glytot ), either by acid hydrolysis
(AC) or by enzymatic hydrolysis (EZ). As well, it has been
determined that rodent muscle contains two pools of glycogen, macroglycogen (MG) and proglycogen (PG). This MG/PG
determination of Glytot has never been compared with AC or
EZ methods, nor has it been determined whether the two
pools exist in human skeletal muscle. A detailed comparison
of the three methods was performed by using both rodent and
human muscle. It was found that repeated analysis of independent portions of muscle generally gave coefficients of variation of ,10%. The PG fraction was always in excess of MG,
which was 6–10% of Glytot in rodent muscle and in human
samples when Glytot was low but increased to ,40% when
Glytot was high. It was found that AC and EZ Glytot were not
statistically different (P , 0.05), nor was there a difference
between the MG1PG Glytot and that determined by AC or EZ.
The Glytot from MG1PG extraction had a strong correlation
with the values obtained by either AC (r 5 1.0) or EZ (r 5
0.96). These data suggest that MG1PG do exist in human
skeletal muscle and can be measured reliably in biopsy-sized
samples. All three methods give an accurate representation of
human Glytot and are comparable in their precision.
MACROGLYCOGEN AND PROGLYCOGEN IN HUMAN SKELETAL MUSCLE
Furthermore, PG and MG concentrations have not
been examined in human muscle under any circumstances. However, before studies into the physiological
response and regulation of these pools can be undertaken, a thorough evaluation of these methods is
required. The purposes of this study were 1) to compare
AC and EZ determinations for total glycogen; 2) to
establish whether MG and PG exist in human muscle
and whether they can be measured in a ‘‘biopsy-sized’’
sample; 3) to evaluate the reproducibility of the AC, EZ,
PG, and MG determinations; and, finally, 4) to compare
the AC, EZ, and MG1PG values for total muscle
glycogen.
METHODS
tracted from the total concentration determined for the
exogenous plus muscle sample. To find the percent recovery,
this value was divided by the exogenous glycogen concentration and then multiplied by 100.
Analysis. The AC glycogen method used was adopted from
Passonneau and Lauderdale (15) and is described as follows.
Freeze-dried muscle samples of between 2 and 3 mg were
hydrolyzed with 2 M HCl and then were heated for 2 h at
85–90°C, followed by neutralization with 2 M NaOH. The
extracts were then analyzed in duplicate fluorometrically by
using the Bergmeyer (3) method for determining glucosyl
units.
The EZ method used was the amyloglucosidase method; 0.1
M NaOH was added to 2–3 mg dry muscle, and samples were
incubated for 10 min at 80°C to destroy ‘‘background’’ glucose
and hexose monophosphates. Then, the samples were neutralized by a combination of 0.1 M HCl, 0.2 M citric acid, and 0.2
M Na2HPO4. Amyloglucosidase was added, and samples were
incubated for 1 h at room temperature while glycogen degradation took place. The samples were analyzed in duplicate
spectrophotometrically at 340 nm by using the method of
Passonneau and Lowry (16).
The method for determination of PG and MG fractions was
based on that described by Alonzo et al. (1) and Lomako et al.
(12, 14) and is also similar to that described by Jansson (10)
for acid-soluble, and -insoluble, glycogen. Ice-cooled 1.5 M
PCA (200 ul) was added to 1.5–3 mg of freeze-dried muscle
samples in 5-ml Pyrex tubes. The muscle was pressed against
the glass tubes with a plastic rod to ensure that all the muscle
was exposed to acid. The extraction continued on ice for 20
min. The samples were centrifuged at 3,000 revolutions/min
for 15 min, after which 100 µl of the PCA supernatant were
removed, placed in Pyrex tubes, and used for the determination of MG. The remaining PCA was discarded, and the pellet
was kept for the determination of PG. One milliliter of 1 M
HCl was added to the MG and to the PG sample; the former
was vortexed, whereas the pellet of the latter was pressed
against the glass with a plastic rod. The tube weights were
then recorded. The tubes were sealed with fitted glass stoppers, and all of the samples were placed in the water bath
(100°C) for 2 h, after which the tubes were reweighed and any
change of .50 µl was rectified with the addition of deionized
water. The samples were then neutralized with 2 M Trizma
base, vortexed, centrifuged at 3,000 revolutions/min for 5
min, and transferred to Eppendorf tubes for analysis of
glucosyl units by using the method of Bergmeyer (3) or stored
at 280°C. Subsequent determination (within 1 wk) of muscle
glycogen in frozen MG or PG extract gave the same values as
fresh extract.
Statistics. The total glycogen for the MG/PG determination
was obtained by summing the two fractions. The fraction of
MG to PG was determined by dividing the concentration of
the MG by the summed total (MG/MG1PG) and reporting it
as a percent (3100%). The coefficient of variation (CV) was
obtained by the formula SD/mean. Linear regression analysis
was performed within comparison sections, and 95% confidence intervals were used to determine agreement among
analysis methods. T-tests were used to determine difference
between means of total glycogen for each of the three methods. In all comparisons, differences were accepted as significant at the 0.05 probability level.
RESULTS
Comparison I. The important findings of this section
were that the three measurements were reproducible
in rat muscle and that MG1PG was reliable for biopsy-
Downloaded from http://jap.physiology.org/ by 10.220.33.5 on June 18, 2017
To evaluate the precision and variability and compare the
various methods (AC, EZ, and MG1PG separation) of measuring muscle glycogen, a variety of approaches was employed.
The muscle samples had been collected during a number of
different studies and conditions ranging from exhaustive
exercise to resting muscle. Every sample in each comparison
weighed 1–3 mg dw (i.e., a biopsy-sized sample) and was
measured in duplicate.
Comparison I. The first comparison used rodent muscle
because most of the initial MG/PG data are derived from this
species and because large muscle samples could be obtained.
Muscle samples of various known glycogen concentrations
from rodent hindlimb were freeze-dried, combined, and mixed
thoroughly to create three large pools for repeated measures.
A total of 45 samples was analyzed. Independent samples
(n 5 15) from each pool were measured for glycogen by using
AC (n 5 5), EZ (n 5 5), or MG1PG (n 5 5).
Comparison II. Human muscle has a much wider range of
muscle glycogen concentration than does rodent muscle.
Hence human muscle samples, which from previous determination were known to have a wide range of total glycogen
concentration, were combined and mixed thoroughly to give
three pools of distinctly different glycogen concentrations. A
total of 18 samples was analyzed for this comparison. Independent samples (n 5 6) from each pool were measured for
glycogen by using AC (n 5 3) or MG1PG (n 5 3).
Comparison III. Individual human muscle samples were
analyzed by using each of the three methods. These were
studied to compare the AC, EZ, and MG/PG determinations
when total glycogen concentration is measured in small
biopsy samples. Because of limited sized, not all samples
could be analyzed by using all three methods. MG/PG was
performed on every sample (n 5 50), EZ on 43 samples, AC on
20 samples, and, of the latter, 15 were analyzed by EZ as well.
Comparison IV. Human muscle samples (n 5 28) were
obtained from an independent laboratory, in which they had
previously been analyzed for total glycogen by EZ. These were
subsequently measured for MG and PG by a technician blind
to the existing data for EZ and then were compared with the
prior independent determinations.
Comparison V. Rat muscle was used to determine the
percent recovery of a given amount of exogenous glycogen by
using MG1PG separation. A single pool of muscle was used,
and a portion was analyzed for MG1PG. A glycogen solution
was made from oyster glycogen (Sigma G-8571), and a portion
of this solution was analyzed for glucosyl units. Muscle
samples had a known quantity of this exogenous oyster
glycogen solution added during the PCA-extraction procedure
(n 5 9). After the samples were assayed for glucosyl units, the
previously determined MG1PG concentration was sub-
909
910
MACROGLYCOGEN AND PROGLYCOGEN IN HUMAN SKELETAL MUSCLE
Table 2. Comparison of MG1PG and AC
determinations in 3 pools of human
skeletal muscle glycogen
Measurement
MG
Low
Medium
High
PG
Low
Medium
High
MG1PG
Low
Medium
High
AC
Low
Medium
High
Mean
SD
SE
CV
5.8
34.1
85.7
0.2
7.2
5.4
0.1
4.1
3.1
3.7
21.0
6.3
38.1
147.7
254.5
0.1
10.1
15.2
0.03
5.9
8.7
0.2
6.9
6.0
43.8
181.8
340.2
0.2
11.5
10.2
0.1
6.7
5.9
0.5
6.3
3.0
48.8
198.5
350.2
6.9
4.8
12.6
4.0
2.7
7.3
14.0
2.4
3.6
All values are mmol/kg dw, except CV, which is reported as a
percent; n 5 3, with each determination performed in duplicate.
0.99, and 0.97, respectively. The ratio of MG to PG
varied greatly in these individual samples, with MG
representing from 6 to 38% depending on total glycogen
concentration. The percentage of MG relative to PG is
not constant. With increases in total glycogen (MG1PG),
the percentage of MG increases as well. The relationship is curvilinear (Fig. 5). There appears to be a cluster
of MG concentrations ranging from 4 to ,50 mmol
glucosyl units/kg dw when the total glycogen concentration is ,300. At higher glycogen concentrations, the PG
concentration does not increase a great deal, but the
Table 1. Comparison of EZ, AC, and MG/PG
concentrations in 3 pools of rodent muscle glycogen
Measurement
MG
Low
Normal
Normal
PG
Low
Normal
Normal
MG1PG
Low
Normal
Normal
EZ
Low
Normal
Normal
AC
Low
Normal
Normal
Mean
SD
SE
CV
3.9
11.7
12.6
0.9
1.6
1.4
0.4
0.7
0.6
22.0
13.3
10.7
54.1
128.6
116.6
3.3
14.9
12.4
1.5
6.7
5.5
6.1
11.6
10.6
58.0
138.8
129.2
4.0
15.8
12.9
1.8
7.1
5.8
6.9
11.4
10.0
58.3
110.9
121.3
4.3
4.0
3.7
1.9
1.8
1.7
7.4
3.6
3.1
53.9
114.4
122.8
6.3
14.3
29.6
2.8
6.4
13.2
11.6
12.5
24.1
All values are in mmol/kg dry weight (dw), except for coefficient of
variation (CV), which is reported as a percent; n 5 5, with each
determination performed in duplicate. EZ, enzymatic hydrolysis; AC,
acid hydrolysis; MG, macroglycogen; PG, proglycogen.
Fig. 1. A comparison of total muscle glycogen in 3 pools of human
muscle by using acid hydrolysis (AC) and macroglycogen
(MG)1proglycogen (PG) methods (comparison II) Each point is a
single sample from 1 of the 3 pools of muscle. dw, Dry wt. Solid line,
linear regression analysis: y 5 1.01x 1 8.38, r 5 1.0, n 5 9; dashed line,
line of identity with slope 5 1; dotted lines, 95% confidence interval.
Downloaded from http://jap.physiology.org/ by 10.220.33.5 on June 18, 2017
sized samples. Generally, CVs were within the published range, although the CV increased with lower
total glycogen concentration (Table 1). However, SE
values were small, and the higher CV was a reflection
of the low glycogen concentration. One of the ‘‘normal’’
glycogen pools had a high CV for AC because the value
for one of the five samples was unusually high.
In a comparison of the AC and EZ methods, a
positive, linear relationship was found (r 5 1.0), and
there was no significant difference between the values
determined by the two methods. Values from the
MG1PG determination were not significantly different
from those from either the EZ or the AC methods.
Comparison II. In the pooled human muscle, the
repeatability was similar to that in comparison I (Table
2). Over a wide range of total glycogen concentrations,
the SD and CV of the MG/PG determination in human
muscle were within the range reported by Harris et al.
(6). The MG/PG determination demonstrated a strong
correlation with the AC method (r 5 1.0), and the line of
identity with a slope equal to 1 fell within the 95%
confidence interval, illustrating that there was no
difference between the two methods. (Fig. 1) There was
no significant difference in the total glycogen values
determined by AC and by MG1PG.
Comparison III. The third comparison of muscle
glycogen for individual biopsy samples of human muscle
presents evidence that the MG/PG determination is
comparable to the AC and EZ methods (Figs. 2–4). The
correlation coefficients as well as the line of identity
and regression line not being significantly different
demonstrate how similar the three methods were in
giving a precise measurement of total glycogen. The
correlation coefficients for EZ vs. MG1PG (n 5 43), AC
vs. MG1PG (n 5 20), and EZ vs. AC (n 5 15) are 0.96,
MACROGLYCOGEN AND PROGLYCOGEN IN HUMAN SKELETAL MUSCLE
MG fraction becomes a greater contributor, although
never the equal of the PG fraction.
Comparison IV. In this comparison human biopsy
samples were analyzed for MG/PG and then compared
with EZ determinations made by an independent laboratory. Again, the MG1PG total glycogen correlated
well (r 5 0.97) with the values previously measured by
Fig. 3. A comparison of total muscle glycogen in human muscle by
using enzymatic (EZ) and MG1PG methods (comparison III). Points
and lines are defined as in Fig. 1. Linear regression analysis: y 5
0.84x 1 30.0, r 5 0.96, n 5 43.
Fig. 4. A comparison of total muscle glycogen in human muscle by
using EZ and AC methods (comparison III). Points and lines are
defined as in Fig. 1. Linear regression analysis: y 5 0.87x 1 14.6, r 5
0.97, n 5 15.
using the EZ method (Fig. 6). Again, there was no
significant difference between the line of identity and
regression line.
Comparison V. This part of the study was designed to
test how much of a given amount of glycogen is actually
recovered by using the MG and PG extraction procedure. A 10-µl sample of a 43 µM solution of exogenous
glycogen was added to a measured amount of rat
muscle, and the glycogen concentration was compared
with the concentration of an independent aliquot from
the same pool. The percent recovery averaged 105 6
8.5% (n 5 9), and the exogenous glycogen was entirely
recovered in the MG portion.
Fig. 5. A comparison of MG values and MG1PG total values
obtained in n 5 50 individual muscle samples.
Downloaded from http://jap.physiology.org/ by 10.220.33.5 on June 18, 2017
Fig. 2. A comparison of total muscle glycogen in human muscle by
using AC and MG1PG methods (comparison III). Points and lines
are defined as in Fig. 1. Linear regression analysis: y 5 0.92x 1 3.8,
r 5 0.99, n 5 20.
911
912
MACROGLYCOGEN AND PROGLYCOGEN IN HUMAN SKELETAL MUSCLE
DISCUSSION
This study was designed to compare the two wellaccepted methods of measuring the total glycogen in a
muscle biopsy sample over a wide range of total glycogen concentrations. It was also conducted to determine
whether MG and PG can be measured in biopsy-sized
samples and whether this method can be applied to
human tissue. The precision of this new technique was
also evaluated and compared with that of the two
accepted methods. This study presents the first determinations of MG1PG for human muscle and strongly
suggests that there are two forms, and probably two
metabolic pools, of glycogen in human muscle.
In rat muscle (comparison I), there was no statistical
significance in total glycogen between the AC and EZ
methods. In addition, the AC and EZ methods were
found to be equivalent in reproducibility (Table 1).
Similarly, in human muscle we found that there was no
systematic difference in the two measurements because
the line of identity with a slope of 1 was within the
limits of the 95% confidence interval (Fig. 1). This is in
agreement with a comment made by Jansson (10).
Harris and Hultman (6) determined that in the measurement of muscle glycogen, there was a variation of
8.43 mmol glucosyl units/kg dw because of analytic
procedure error alone. The SD for routine analysis of
muscle glycogen samples taken during exercise experiments was found to be between 10 and 13 mmol
glucosyl units/kg dw (6). Our data for rodent and
human muscle glycogen are in agreement with these
findings. Therefore, each of the three methods gives low
and similar CV values that are comparable with findings in the literature.
The reproducibility in the pooled human muscle
samples is in agreement with other studies, and the
Downloaded from http://jap.physiology.org/ by 10.220.33.5 on June 18, 2017
Fig. 6. A comparison of total glycogen in human muscle by using EZ
values previously obtained by an independent laboratory and MG1PG
methods (comparison IV). Points and lines are defined as in Fig. 1.
Linear regression analysis: y 5 0.93x 1 19.2, r 5 0.97, n 5 26.
variances (Table 2) in the present study were within the
published range (6). The repeated measurement of MG,
PG, and MG1PG in human muscle (n 5 9) had a CV
range from 0.2 to 21%. The majority of the CV values
displayed in Table 2 lie between 3 and 6%. Essen and
Henriksson (4) found that there was a CV of ,6.3%
when they measured glycogen within single muscle
fibers. In a comparison of the human muscle total
glycogen for the MG/PG determination for the low,
medium, or high concentrations with those determined
by the AC method, there was no statistical significance
among the values. The correlation among these methods was 1.0 with slope of 1, and the line of identity was
almost identical to the regression line, demonstrating
that the MG1PG technique is as precise as the traditional AC and EZ methods (Fig. 1). The MG/PG determination for individual muscle samples (n 5 20) compared with AC also gave extremely high correlations.
Thus the three methods are comparable.
MG/PG values calculated from comparison II suggest
that as the total glycogen concentration rises so does
the percentage of MG. We found that at a MG1PG
concentration of 43.8 mmol glucosyl units/kg dw the
MG/PG was 13:87. As the total increased to 181.8 mmol
glucosyl units/kg dw, the ratio increased to 19:81 and to
25:75 as the total concentration reached 340.2 mmol
glucosyl units/kg dw. The percentage of MG in the
high-concentration pool was significantly higher than
the percentage in the low pool. This is in accordance
with Jansson’s (10) results, which displayed an increasing percentage of soluble glycogen as the total concentration increased. She found that PCA-soluble muscle
glycogen constituted 25% of the total glycogen content,
which was #350 mmol/kg dw, and increases in total
glycogen above that concentration seemed to be mainly
due to the soluble type (MG). In the present study, in
the individual muscle sample with the highest total
glycogen (529.6 mmol glucosyl units/kg dw), the ratio
was 38:62. Alonzo et al. (1) also showed that a greater
percentage of PG relative to MG (molar percentage:
85% PG to 15% MG) in resting rabbit muscle. Interestingly, Friden et al. (5) reported that two populations of
glycogen particles were clearly distinguished and, of
the 144 particles counted in a pool in the intermyofibrillar space, ,76% were of the smaller particle size. This
consensus suggests that MG1PG can be detected by
electron microscopy.
Glucose analyses in the PCA extracts of human
muscle before hydrolysis yielded low values (0.11–3.61
mmol/kg dw) (10). This demonstrated that glycogen
was not hydrolyzed into glucose residues during the
PCA-extraction procedure. Jansson (10) also demonstrated that the relationship between PCA-soluble and
-insoluble glycogen was not influenced by strength of
the PCA in the range from 0.5 to 3 M or the type of acid
(PCA vs. trichloracetic acid). Nor was it affected by the
freeze-dry procedure or the weight of the samples in the
range from 0.2 to 2 mg. The benefit of using the
MG1PG determination is in the ability to separate the
two fractions of muscle glycogen and hence to study the
metabolism of each one individually.
MACROGLYCOGEN AND PROGLYCOGEN IN HUMAN SKELETAL MUSCLE
The authors thank Premila Sathasivam for technical assistance
and Dr. Mark Tarnopolsky for the contribution of muscle samples for
comparison IV.
This study was supported by the Natural Sciences and Engineering
Research Council (NSERC) of Canada and a NSERC Studentship in
collaboration with the Gatorade Sports Science Institute (K. B. Adamo).
Address for reprint requests: T. Graham, Human Biology and
Nutritional Sciences, Univ. of Guelph, Guelph, Ontario, Canada N1G
2W1 (E-mail: [email protected]).
Received 30 June 1997; accepted in final form 6 November 1997.
REFERENCES
1. Alonzo, M., J. Lomako, W. Lomako, and W. Whelan. A new
look at the biogenesis of glycogen. FASEB J. 9: 1126–1137, 1995.
2. Beck-Nielsen, H., A. Vaag, P. Damsbo, A. Handberg, O.
Nielsen, J. Henriksen, and P. Thye-Ronn. Insulin resistance
in skeletal muscle in patients with NIDDM. Diabetes Care 15:
481–428, 1992.
3. Bergmeyer, H. Methods of Enzymatic Analysis New York:
Academic, 1974, vol. 3, p. 1128–1131.
4. Essen, B., and J. Henriksson. Glycogen content of individual
muscle fibres in man. Acta Physiol. Scand. 90: 645–647, 1974.
5. Friden, J., J. Seger, and B. Ekblom. Topographical localization of muscle glycogen: an ultrahistochemical study in the
human vastus lateralis. Acta Physiol. Scand. 135: 381–391,
1989.
6. Harris, R., E. Hultman, and L.-O. Nordesjo. Glycogen, glycolytic intermediates and high energy phosphate determination in
biopsy samples of musculus quadriceps femoris of man at rest.
Methods and variance of values. Scand. J. Clin. Lab. Invest. 33:
109–120, 1974.
7. Hermansen, L., and O. Vaage. Lactate disappearance and
glycogen synthesis in human muscle after maximal exercise.
Am. J. Physiol. 233 (Endocrinol. Metab. Gastrointest. Physiol. 2):
E422–E429, 1977.
8. Huang, M., C. Lee, R. Lin, and R. Chen. The exchange
between proglycogen and macroglycogen and the metabolic role
of the protein-rich glycogen in rat skeletal muscle. J. Clin. Invest.
99: 501–505, 1997.
9. Hultman, E. Muscle glycogen in man determined in needle
biopsy specimens. Method and normal values. Scand. J. Clin.
Lab. Invest. 19: 209–217, 1967.
10. Jansson, E. Acid soluble and insoluble glycogen in human
skeletal muscles. Acta Physiol. Scand. 113: 337–340, 1981.
11. Karlsson, J. Lactate and phosphagen concentrations in working
muscle of man (Abstract). Acta Physiol. Scand. Suppl. 358: 81,
1971.
12. Lomako, J., W. Lomako, and W. Whelan. Proglycogen: a
low-molecular-weight form of muscle glycogen. FEBS Lett. 279:
223–228, 1991.
13. Lomako, J., W. Lomako, and W. Whelan. A self-glucosylating
protein is the primer for rabbit muscle glycogen biosynthesis.
FASEB J. 2: 3097–3103, 1988.
14. Lomako, J., W. Lomko, W. Whelan, R. Dombro, J. Neary,
and M. Norenberg. Glycogen synthesis in the astrocyte: from
glycogenin to proglycogen to glycogen. FASEB J. 7: 1386–1393,
1993.
15. Passonneau, J., and V. Lauderdale. A comparison of 3 methods of glycogen measurement in tissues. Anal. Biochem. 60:
405–412, 1974.
16. Passonneau, J., and O. Lowry. Enzymatic Analysis: A Practical Guide. Clifton, NJ: Humana, 1993, p. 177–178.
17. Roe, J., I. Bailey, R. Gray, and I. Robinsson. Complete
removal of glycogen from tissues by extraction with cold TCA
acid solution. J. Biol. Chem. 236: 1244–1246, 1961.
18. Sabina, R., J. Swain, W. Bradley, and E. Holmes. Quantitation of metabolites in human skeletal muscle during rest and
exercise: a comparison of methods. Muscle Nerve 7: 77–82, 1984.
19. Stetten, M., H. Katzen, and D. Stetton. A comparison of the
glycogen isolated by acid and alkaline procedure. J. Biol. Chem.
232: 475–488, 1958.
20. Stetten, D., and M. Stetten. Glycogen metabolism. Physiol.
Rev. 40: 505–537, 1960.
21. Willstaetter, R., and M. Rhodewald. Protein binding of physiologically important substances. The condition of glycogen in
liver, muscle and leukocytes. Z. Physiol. Chem. 225: 103–124,
1934.
Downloaded from http://jap.physiology.org/ by 10.220.33.5 on June 18, 2017
There are reports, with regard to different pools of
glycogen, that date back to 1900, when Nerking (20),
and later, to 1934, when Willstaetter and Rhodewald
(21) concluded that tissue glycogen appeared in two
forms: an acid-extractable or free form and an acidnonextractable or protein-fixed form. It was not resolved whether the glycogen-protein complexes were
artifacts or whether they constituted a physiological
entity. Since then, investigators have shown different
sizes, different concentrations, and different solubilities of glycogen molecules. Integrating the results of
past studies suggests that data have consistently
pointed to the existence of two different pools of muscle
glycogen, with the smaller molecular/granular form
being most common.
Lomako et al. (14) were able to demonstrate the
movement of labeled glucose initially into the PG
fraction, and then 30 min later the label appeared in
the MG fraction while simultaneously falling in the PG
fraction. A similar study conducted by Huang et al. (8)
in resting rodent muscle demonstrated the incorporation of [3H]glucose into the PG fraction first and then
into the MG fraction under conditions of moderate
increases in [3H]glycogen synthesis. These results suggest that PG is the plausible precursor and that PG
synthesis precedes that of MG. From the limited human data in the present study, it is speculated that PG
is the small and dynamic, intermediate form of muscle
glycogen, whereas MG is the larger storage form that
appears to increase on a relative basis as the total
glycogen increases. It is possible that when the glucose
environment is favorable and the PG has reached a
critical limit, a portion is synthesized into MG, and it is
conceivable that the two pools may eventually equilibrate. With this new separation technique, it is now
possible to study each pool of glycogen exclusively to
understand how the fractions change under conditions
of breakdown and resynthesis.
In summary, the AC and EZ determinations are not
systematically different. MG and PG do exist in human
skeletal muscle and can be measured accurately in
biopsy-sized samples. The AC, EZ, and MG1PG determinations are reproducible and give the same values
for total muscle glycogen.
913