PROGLYCOGEN AND MACROGLYCOGEN IN EQUINE SKELETAL MUSCLE
A Thesis Presented to The Faculty of Graduate Studies of The University of Guelph
by JOHAN BROJER
In partial fulfillment of requirements for the degree of Master of Science April, 2001
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PROGLYCOGEN ANli) MACROGLYCOGEN IN EQULNE SKELETAL MUSCLE
Johan Brojer Advisor: uni ver si^ of Guelph, 2001 Dr. Henry R. St-pfli
The objectives of this thesis were to investigate the existence of proglycogen (PG) and macroglycogen (MG) in equine skeletal muscle; to compare the analysis of total glycogen by two methods (acid hydrolysis and MGtPG determination) for total muscle glycogen fiom biopsy sized sarnples and to evaluate the effect of extraction time and perchloric acid (PCA) concentration on the recovery of MG and PG. The first study indicated that extraction time had very little effect on the recovery of PG and MG but there was an effect of PCA concentration. Extraction with 0.5 and 1.5 M PCA yielded similar values for PG and MG but extraction in 3.0 M PCA yieIded lower PG and higher
MG concentrations (W0.05). The results f?om the second study showed that PG and MG exist in equine skeletal muscle and the measurement techniques for total glycogen analyzed by the two methods were very comparable. The present study was conducted at the Departrnent of Clinical Studies - Large Animal Interna1 Medicine, Ontario Veterinary College, University of Guelph, Canada.
These studies were generously supported by the EP Taylor Equine Trust Fund.
During the the spent completing this work, nurnerous people have been involved, and 1 would like to express my thanks to au who have contributed in various ways.
1wish to express my sincere gratitude to:
Dr. Henry Stampfli, my supervisor, for introducing me to the interesting field of muscle physiology and for stirnulating my interest in large animal interna1 medicine. You have been such a nice person to work with - both in the large animal chic and in the area of research. Thank you for your guidance, patience and your unfailing encouragement and interest and for aIways keeping me cheerfül when progress was slow. 1 consider you and your family as very good fnends and 1 am very happy for a11 the dimers you provided me and my wife during our time in Canada.
The members of my research cornmittee: Dr. George Heigenhauser and Dr. Tim Lumsden, for your constructive criticism on the manuscript, for interesting scientific discussions during the cornmittee meetings and for offerhg valuable advice conceming the thesis.
Dr. Terry Graham, rny CO-author,for your excellent guidance in the field of pro- and macroglycogen and for your always optirnistic and friendly attitude. Your constructive criticism and comments on the manuscripts have always been appreciated. Thanks a lot for letting me have access to your research Iaboratory.
The owners of the valuable endurance ride horses, for giving me the opportunity to study their fine horses.
Dr. Birgitta Essén-Gustavsson for your knowledge, enthusiasm, support and for al1 interesting scientific discussions. To write a thesis having the advisors and CO-authoron the other side of the Atlantic sea does not make things easier. Therefore, 1 am gratefb! for a11 your help - to have a person at the same department who 1could ask questions to right away.
Premila Sathasivam for al1 your help, support and knowledge. You had al1 the answers 1 needed when prob1ems suddenly appeared in the laboratory. Kristi Adamo and Jane Shearer for introducing me to the analytical methods of pro- and macrogfycogen.
My colleagues at the department of Equine Interna1 Medicine, Swedish Veterinary College, Susanne Demmers, Pia Funkquist, Agneta Gustafsson, John Pringle, Miia Riihimaki and Katarina Schuback for your help with the schedule so 1 finally got the time off for writing the thesis,
Kim Kultiman for a11 your knowledge about statistics and your help with the computer program Minitab.
Caroline, my wife and very best fiend, for being the most important person in my life and for sharing rny interest in hunting and the outdoors. Thank you for your love, help, support and encouragement but most of al1 for being my wife.
Rebecka, my newborn daughter, for helping me remember what the important things are in life.
My four-footed fiends Smaragd and Amethist (my horses) for sharing al1 those nice moments in the countryside and Siska (my dog) for being a nice hunting partner but also for being a Company during my thesis-writing at night. TABLE OF CONTENTS
TABLE OF CONTENTS...... iii
DECLARATION OF WOEX PERFORMED ...... v
LIST OF TABLES ...... vi ... LIST OF FIGURES ...... vlzl
CHAPTER 1
1. 0 General Introduction ...... -1 1.1 Statement of Goals and Hypotheses...... 5
1.2 General Literature Review
1.2.1 Introduction ...... -7 1.2.2 Glycogen ...... 7 1.2.3 Glycogenin ...... -8 1.2.4 Proglycogen and macroglycogen ...... 9 1.2.5 Theoretical modefing of the glycogen structure...... 12 1.2.6 Storage of glycogen in skeletal muscle - the relationship between pro- and macroglycogen ...... -13 1.2.7 Synthesis and glycogenolysis of pro- and macroglycogen...... -15 1.2.8 Reference list ...... 23
CHAPTER 2 The effect of extraction time and perchloric acid concentration on the recovery of pro- and macroglycogen from muscle biopsies from the horse
2.0 Abstract ...... 30 2.1 Introduction ...... 30 2.2 Material and Methods ...... 32 2.3 Results ...... 35 2.4 Discussion ...... 37 2.5 Acknowledgments...... 40 2.6 References ...... -45
iii
DECLARATION OF WORK PERFORMED
I declare that al1 work reported in this thesis was performed by me. Biopsy sarnpling procedures were undertaken with the assistance of Dr. Henry R, Stiimpfli. LIST OF TABLES TABLE Theoretical increase in glycogen levels above basal values...... 22
Mean pro-, macro- and total glycogen concentrations for different extraction times...... 41
Mean proglycogen concentration for different combinations of extraction tirne and perchloric acid concentration...... -42
Mean rnacroglycogen concentration for different combinations of extraction time and perchloric acid concentration...... 43
Mean total glycogen concentration for different combinations of extraction time and perchloric acid concentration...... --. -44
Reproducibily for total glycogen in duplicate analyses from the same muscle biopsy for the acid hydrolysis and the pro- and macroglycogen technique.. -65
Reproducibilty in duplicate biopsies obtained at the sarne site in the glzlteza medius muscle...... 66
The effect of sarnpling dep th in the glutetcs medius muscle...... -66
Individual data for proglycogen (PG), macroglycogen (MG), PWMG and acid hydrolysis (AC) fiom 45 individual muscle biopsies...... 94
Individual data for proglycogen (PG), macroglycogen (MG) and PG+MG £kom biopsies obtained at different sarnpling depths...... 96
Individual data for proglycogen (PG), macroglycogen (MG) and PG+MG fkom repeated biopsies obtained fkorn the same site in the middle gluteal muscle.. -97
Individual data for proglycogen (PG),macroglycogen (MG) and PGcMG for different extraction times...... 98
Individuai data for proglycogen for different combinations of exaaction tirnes and perchlo ric acid (PCA) concentrations...... 99
Individual data for macroglycogen for different combinations of extraction times and perchloric acid @CA) concentrations...... -10 1 5.7 Individual data for proglycogen + macroglycogen for different combinations of extraction times and perchloric acid (PCA) concentrations...... 103
5.8 Individual data for glucose concentration in aliquots fiom the macroglycogen fiactions before hydrolysis in HC1 ...... -105
vii LIST OF FIGURES
1.1 Schematic structure of two outer branches of a glycogen particle with a(1-4) and a(1-6) linkages...... 19
1.2 Glycogen synthesis on a protein core as proposed by Krisman and Barengo...... -...... -19
1.3 The interaction between glycogenin and glycogen synthase in the synthesis of glycogen...... 20
2 -4 Mode1 of glycogen synthesis as proposed by Whelan and colleagues...... 21
3.1 Linear regression of total glycogen measured by acid hydrolysis and pro- and macroglycogen determinatron..*...... -62
3.2 Scatter diagram of the paired differences between methods for analysis of total glycogen (pro- and macroglycogen determination and acid hydrolysis)...... 63
3.3 A comparison of macroglycogen concentrations and total glycogen concentration obtained kom individual muscle biopsies (n=3 1) from the middle gluteal muscle...... 64
viii CHAPTER 1
1.0 GENERAL INTRODUCTION
The metabolisrn of muscle glycogen has been intensely studied in both human and equine exercise physiology. The introduction of the percutaneous muscle biopsy technique by Bergstrom has enabled our understanding of both equine and hurnan muscle to expand tremendously over the last 30-40 years. The most fiequently exarnined muscle in the l-iorse is the middle gtuteal.
Earlier biochemical (Kits Van Heijningen and Kemp 1955, Stetten Jr. and Stetten
1960, Jansson 1981) and electron rnicroscopic studies (Friden et al. 1989) indicated that glycogen was not a uniform molecule. Recent studies have identified two pools of glycogen in rat and human skeletal muscle; proglycogen (PG) and macroglycogen (MG)
(Lomako et al. 1993, Alonso et al. 1995b, Adamo and Graham 1998). Lomako, Whelan and coworkers (Lomako et al. 1993, Alonso et al. 1995b) were the first to describe these two forms of glycogen in detail. Proglycogen is the srnaller molecule with a molecular weight of up to 400 kDa whereas MG is the larger molecule, which can reach a rnolecular rnass of 10,000 kDa (Lornako et al. 1991)- Both molecular forrns contain identical arnounts of protein but different amounts of associated carbohydrate. Proglycogen is made up of approxirnately 10% protein whereas macroglycogen contains only approximately
0.4%. Since the two forms have different ratios between protein and carbohydrate they cm be separated on the basis of their solubility in acid. Proglycogen precipitates in tnchloroacetic acid (TCA) and perchloric acid (PCA) whereas MG is soluble in these acids (Lomako et al. 1991, Adamo and Graham 1998). Adamo and Graham (1998) have recently validated a separation technique for PG and MG for hurnan and rat skeletal
muscle where freeze dried muscle biopsies were exposed to ice-cooled 1.5 M PCA for 20
minutes. Jansson (1981) showed earlier that the measured ratio between the two pools of
glycogen in human skeletal muscle was not influenced by the type of acid (PCA vs. TCA)
or by the strength of the PCA in the range between 0.5 and 3 M. Although a number of
studies have used the separation technique for PG and MG in rat and human muscle there
have been no reports showing the stability of the two fractions with extraction the. In
case the concentration of PG and MG changes with extraction time without reaching a
stable level the separation technique must be questioned.
The resting rat skeletal muscle contains approximately 88% of the total glycogen in
the PG form (Hansen et al. 2000) whereas resting human skeletal muscle contains 66-77%
in the PG form(Graham et al. 2001). In exercising human skeletal muscle the net rate of
glycogenolysis is almost always greater for the PG pool compared to the MG pool
(Graham et al. 2001). In the rat skeletal muscle however, the relative utilization during electrical stimulation is greater for the MG pool (Derave et al. 2000). These findings indicate that there are species differences with regard to both metabolism and relationship between PG and MG in the skeletal muscle. The separation technique for PG and MG therefore requires fiirther validation for the equine skeletal muscle before studies of the physiological response of glycogenolysis, synthesis and regulation of the two forrns cm be conducted experimentally in exercising horses.
The equine muscle, as for most other mammalian muscles, consist of a mosaic of muscle fiber types with varying metabolic and contractile properties. In the skeletal muscle the glycogen content varies between muscle fibers types. The type II fibers have higher glycolytic capacity and higher glycogen content compared to the type 1 fibers
(Snow et al. 198 1, Valberg 1986). During exercise the number and types of fibers recruited varies within the muscle depending on the type, speed and duration of exercise
(Essen-Gustavsson et al. 1984, Valberg 1986, White and Snow 1987, Gottlieb 1989). This variation in fiber recruitment during exercise results in differences in the glycogen depletion pattern. The non-uniforrn distribution of glycogen between muscle fiber types and the variation in fiber recruitment during exercise gives the potential for high variability among biopsy samples for both pre- and post exercise biopsy samples. Despite this variation several studies have shown that analysis of glycogen in muscle biopsies from both horses (Lindholm and Piehl 1974, Snow et al. 1981) and man (Hultman 1967,
Essen and Henriksson 1974, Harris et al. 1974) yielded results with Iow variation .
The individual muscle is not uniform in its fiber distribution due to different activity profiles at different levels of the muscle. In most muscles the highest proportion of type II fibers are found superficially with an increasing proportion of type I fibers in the deeper parts (Snow and Valberg 1994). In one study of the fiber type variation within the middle gluteal muscle of the horse there was an increase in the mean percentage of the type 1 fibers from 21 -6% at a sarnpling depth of 2 cm to 61.5% at a sarnpling depth of 8 cm
(Serrano et al. 1996). There was a corresponding decrease in the mean percentage of the type W fibers fiom 42% to 4.5% with increasing sarnpling depth while the mean concentration of the type IIA fibers remained very sirnilar (mean range 34 - 36.5%) between the different sarnpling depths. This non-uniformity of fiber types within the gluteal muscle is an important consideration for biochemical analysis of muscle biopsies in the horse. This study was undertaken to investigate if PG and MG exist in equine skeletal muscle and to validate the separation technique for the two fiactions of glycogen in horse muscle. Total glycogen analyzed fkorn freeze dried muscle biopsies by acid hydrolysis or by the separation technique for PG and MG were cornpared. Furthemore, the effect of different sampling depths in the rniddle gluteal muscle, PCA concentrations and extraction times on the concentration of PG and MG were evaluated. 1.1 STATEMENT OF GOALS AND HYPOTHESES
The overall goal of the studies descnbed in this thesis was to investigate proglycogen and macroglycogen in equine skeletal muscle. The central hypothesis was:
Proglycogen and macroglycogen exist in equine skeletal muscie and can be reliably measured in b iopsy-shed samples.
These studies were divided into two main components; the hypothesis and objectives for each of these components were as follows:
Cliapter2: The effect of different extraction times and perchloric acid
concentrations on the recovery of pro- and macroglycogen from muscle
biopsies from the horse.
Hypo th esis:
1. The PG and MG pools remain constant over a wide range of PCA
concentrations and extraction tirnes.
Objectives:
1. To compare the effect of extraction time in the range between 10 and 120
minutes on the recovery of PG and MG fiom fieeze dried muscle biopsy
sarnples.
2. To compare the effect of the combination of different extraction times
and PCA concentrations on the relationship between PG and MG from
fieeze dried muscle biopsy samples. CItnpter 3: Analysis of proglycogen and rnacroglycogen in muscle biopsies from the
horse.
Hypo th esis:
1. Proglycogen and macroglycogen can be reliably measured in i?eeze ciried
muscle biopsy samples fiom the horse.
Objectives:
2. To compare analysis of total glycogen in muscle biopsies hmhorses by
two methods (AC and PG-MG determination technique).
2. To evaluate the reproducability of the AC, MG and PG determination.
3. To compare the effect of different sampling depths in the gluteal medius
muscle on the concentrations of PG and MG. 1.2 LITERATURE REVIEW
1-2.2 INTRODUCTION
Glycogen is a very large branched polymer of glucose residues that is built upon a
protein core. It serves as a readily mobilized storage form of glucose in most animal ce11
types. The liver and muscle serve as the two main sites of glycogen storage. The liver has
the highest glycogen concentration and this organ has an important role in the
maintenance and regulation of blood glucose. Despite the higher relative glycogen
concentration in the liver the skeletal muscle contains a larger arnount of glycogen due to
its greater rnass. The primav purpose of the glycogen stores in the skeletal muscle is to
provide readily accessible energy for muscle work (Lehninger 1982).
1.2.2 GLYCOGEN
Glycogen consists of glucose residues linked together by a-1,4-glycosidic bonds or
a-l,6-glycosidic bonds. The cc- 1,6-glycosidic links are the branch points of which there
are one in about ten residues whereas the rest of the links between the glucose residues are
created by a-1,4-glycosidic links (Fig 1). The branched structure of glycogen has several
advantages compared to a non-branched structure: 1) it creates a more compact structure,
2) it increases the rate of glycogen synthesis and degradation by increasing the nurnber of terminal residues - the action sites of glycogen synthase and phosphorylase, 3) it increases the solubility of glycogen (Stryer 1988). 1.2.3 GLYCOGENIN
The biosynthesis of glycogen involves both a specific initiation phase and a phase of elongation. The enzyme glycogen synthase (GS) cannot transfer glucosyl residues fiom uridine-diphosphate-glucose (UDP-glc) without a disaccharide already present and an initiating stage is therefore required (Calder 1991). Krisman and Barengo (1975) suggested that there was a protein backbone to which glucosyl units are transferred at the initiation stage by the activity of a "glycogen initiator synthase" and that the further elongation of the glycogen particle was catalyzed by glycogen synthase and branching enzyme (Fig. 2). Later, Whelan and CO-workers(Rodriquez and Whelan 1985) identified a protein that was covalently attached to glycogen via tyrosine residues. This protein backbone of glycogen was named glycogenin (Pitcher et al. 1987). Further work performed by Whelan (Lornako et al. 1988) and Cohen (Pitcher et al. 1987) showed that glycogenin was not only the protein backbone on which glycogen is synthesized but that it also had self-glucosylating activity. Via self-glucosylation glycogenin forms an oligosaccharide primer of 7-1 1 glucosyl units, which then serves as an effective substrate for glycogen synthase (Alonso et al. 1995b) (Fig. 3).
The protein glycogenin in skeletal muscle has a molecular mass of 37 kDa and consists of 332 amino acids (Campbell and Cohen 1989). The discovery by the group of
Cohan (Pitcher et al. 1988) and Whelan (Lomako et al. 1988) that glycogenin had enzymatic activity by catalyzing a self-glucosylation reaction with UDP-glucose as a donor, was a landmark in understanding glycogenin function. Glycogenin is now classified as a hexosyltransferase (EC 2.4.1.186). The self-glucosylation of glycogenin is stimulated by MnL7 and requires UDP-glc as a substrate (Alonso et al. 1995b). The glycogen is attached to the protein glycogenin by a single glucose-1-O-tyrosyl linkage.
The tyrosine (Rodrïquez and Whelan 1985) was identified by protein chemical methods as
tyr-194 (Campbell and Cohen 1989). Alonso and coworkers fond that when tyr was
replaced with phenylalanine @he) the glycogenin would no longer self-glucosylate, which showed the importance of this site for the activity and function as well as for the anchor of glucose (Alonso et al. 1995a). Duhg the formation of an oligosaccharide primer the glucose residues are attached to each other by a-1-4-glycosidic bonds as in glycogen
(Pitcher et al. 1988). Because of the different chemical nature of these linkages behveen protein and the first glucose residue and the rest of the glucose residues, there was initially a debate as to whether a separate glucosylîransferase was needed to mediate the initial step. It seemed unlikely at the time that one enzyme could be capable of catalyzing both carbohydrate-protein and carbohydrate-carbohydrate bonds (Roach and Skurat 1997).
Later research showed, however, that glycogenin indeed was able to catalyze both linkages and the role of glycogenin in the biosynthesis of glycogen is now fairly well established (Lomako et al. 1990, Alonso et al. 1995b). Even though glycogenin is capable of both tyrosine glucosylation and carbohydrate glucosylation it is not clear if there exist two catalytic sites or only one (Alonso et al. 1995b).
1.2.4 PROGLYCOGEN AND MACROGLYCOGEN
In the traditional mode1 of glycogen synthesis glycogenin, glycogen synthase, branching enzyme and the substrate UDP-glucose are sufficient to account for glycogen synthesis. Recently, Lornako et al. (1991) proposed the existence of an additional discrete intermediate form in the synthesis and degradation of glycogen, termed proglycogen (PG). Using fiesh muscle extracts incubated with UDP['4C]glucose and subjected to SDS-
PAGE they identified a well defined band with high molecular weight of approximately
400 kDa (p400) but no bands with smaller molecular weights. The p400 was glycogen-
like and could be broken down to gIycogenin by amylase treatrnent at 37°C (Lomako et al.
1990). Under normal conditions glycogenin does not exist in a fiee, detectable state in skeletal muscle (Lomako et al. 1990, Smythe et al. 1990). Based on these and other findings Lomako, Whelan and Alonso (Lornako et al. 1992, Lornako et al. 1993, Alonso et al. 1995b) proposed the existence of two forms of glycogen; proglycogen and macroglycogen. According to this mode1 the p400 (progfycogen molecule) was suggested to be a stable intermediate in the synthesis of mature glycogen (rnacroglycogen) and under normal conditions never broken down to glycogenin. Lomako, Whelan and coworkers showed that the formation of proglycogen and macroglycogen by the addition of glucosyl units using UDP-glucose as a substrate were both sthulated by glucose-6-phosphate
(G6P), which is characteristic for glycogen synthase (Lomako et al. 1991). However, the
K, (Michaelis constant) for UDP-glucose was three orders of magnitude Iower for the synthesis of proglycogen cornpared to the synthesis of macroglycogen. Furthermore, the formation of macroglycogen but not the formation of proglycogen was inhibited by ammonium ions (Lomako et al. 1991, Lomako et al. 1993). Thus, the reactions responsible for adding glucosyl units in the formation of proglycogen and macroglycogen showed remarkable differences and they therefore suggested that the synthesis of glycogen from glycogenin to mature glycogen via the formation of proglycogen is catalyzed by three different enzyme activities (Fig. 4) (Alonso et al. 1995b). The first step in the glycogen synthesis is the self- glucosylation by glycogenin. The second step is the formation of proglycogen by proglycogen synthase and the final step is the furmation of macroglycogen by macroglycogen synthase. Whether proglycogen synthase and macroglycogen synthase are two different enzymes or the same enzyme phosphorylated differently is at present unknown (Alonso et al. 1995b).
According to the mode1 proposed by Lomako, Whelan and coworkers proglycogen is a smaller intemediate in the synthesis of mature glycogen (macroglycogen) with a molecular weight of approximately 400kDa. Macroglycogen is the larger glycogen rnolecule which cm reach a molecular weight of 10,000 kDa. Both molecular foms contain identical amounts of protein but different amounts of associated carbohydrates.
Therefore, the two forms differ in the relative proportion of protein to carbohydrate.
Proglycogen contains approxirnately 10% protein and is precipitable in 10% trichloroacetic acid (TCA) whereas rnacroglycogen is soluble in TCA due to its very srna11 amount of protein (approximately 0.4%) (Alonso et al. 1995b).
The hypothesis of the existence of two forms of glycogen is not by any means novel.
Authors to earlier studies have also discussed the possibility of two forms of glycogen.
Willstaetter and Rhodewald (1934) demonstrated the existence of two forms of glycogen separable b y TCA solubility; lyo- and desmoglycogen. Lyoglycogen was soluble in TCA and was believed to be protein-fiee whereas desmoglycogen was believed to contain protein and was therefore insoluble in TCA. Stetten et al. (1958) found that fiactionalization of glycogen by extraction in 5% TCA or by alkaline digestion in KOH resulted in glycogen with different mean molecular weights. In 198 1 Jansson reported that there were two portions of glycogen in human skeletal muscle; one was acid soluble in 1.5
M perchlonc acid (PCA) and one was insoluble. Jansson also demonstrated that the relationship between acid soluble and insoluble glycogen in human skeletal muscle was
not influenced by the type of acid (PCA vs. TCA) or by the strength of PCA in the range
between 0.5 and 3 M (Jansson 1981). The theory of the existence of two glycogen pools
was supported in 1989 by Fridén et al. who identified two different sizes of glycogen
particles in human skeletal muscle by using electron microscopy (Fnden et al. 1989).
In 1997 Adarno and Graham validated a separation technique for MG and PG for
human and rat skeletal muscle where fieeze-dried muscle biopsies were extracted for 20
minutes in 1.5 M PCA. The separation technique was based on the findings fiom both the
work performed by Jansson (1981) and Lomako, Whelan and coworkers (Lomako et al.
1993, Alonso et al. 1995b). Total glycogen analyzed by three different methods:
enzyrnatic hydrolysis, acid hydrolysis with 1M HCI and by the separation technique for
pro- and macroglycogen, yielded comparable results with regard to both total glycogen
concentration and repeatability (Adarno and Graham 1998).
1.2.5 THEORETICAL MODELING OF THE GLYCOGEN STRUCTURE
A theoretical model for glycogen structure has been proposed by Goldsmith and
colleagues (Goldsmith et al. 1982). According to this model the glycogen molecule is
built on a protein core and consists of approximately 4200 glucose chains with an average
length of 13 glucose units each. The nurnber of chains increases by a factor 2 for each new
layer in the glycogen molecule. After 12 layers the surface density of the chains has
increased to the level where merbranching is limited and the growth of the molecule is
self-limiting. The calculated molecular weight of this theoretical model of a glycogen particle is 10' Da, which is in nice agreement with the weight of the glycogen B-particle. The ratio of activities between branching enzyme and glycogen synthase account for the control of the glycogen chain kngth. Experimental evidence has shown that the chains of the different levels (including inner and outer chains) of the glycogen particle are, on average, 13 residues long (Meléndez et al. 1997). During the synthesis of the different layers the ratio between the activities of the two enzymes would have to change in order for the length of the chains to remain constant, which is udikely. Meléndez-Hevia et al.
(1997) suggested that the pro- and macroglycogen mode1 could explain the process.
According to this mode1 two different enzymes, proglycogen synthase and macroglycogen spthase, could be responsible for the constant length of the different parts of the glycogen molecule. The proglycogen molecule consists of 7-8 tiers, which creates a molecular weight of approximately 400 kDa whereas the macroglycogen molecule consists of 12 tiers and a molecular weight of approximately 20,000 kDa.
1.2.6 STORAGE OF GLYCOGEN IN SKELETAL MUSCLE - THE
RELATIONSHIP BETWEEN PRO-AND MACROGLYCOGEN
Human and rat muscle glycogen is composed of pro- and macroglycogen and the relationship between these two forms of glycogen is dependent on the totaI glycogen concentration. In muscle with very low total glycogen concentration alrnost al1 glycogen is present as proglycogen. When the total glycogen concentration increases the relative proportion of proglycogen decreases whereas the relative proportion of macroglycogen increases. Despite the decrease in the relative proportion of proglycogen the absolute proglycogen concentration in fact increases (Jansson 1981, Adarno et al. 1998, Hansen et al. 2000). Jansson (1981) dernonstrated that for hurnan skeletal muscle with normal glycogen concentrations (300-350 mm01 glucosyl unitskg dry weight) the acid soluble fraction (macroglycogen pool) comprised approximately 25%. At total glycogen concentrations over 300-350 molglucosyl unitskg dry weight (dw) the increase in total glycogen was predominantly due to increase in the acid soluble pool. Similar results were obtained by Adarno and Graham (1998) in a study performed on human skeletal muscle.
The proglycogen concentration appeared to reach its maxima1 concentration at 250-300 mm01 glucosyl units per kg dw, which corresponded to a total glycogen concentration of approximately 300-350 mm01 glucosyl units per kg dw. Increase in total glycogen over
350 mm01 glucosy1 units per kg dw appeared to be exclusively due to increase in the macrogIycogen pool.
The skeletaI muscle glycogen stores can be increased above the normal concentrations in rat and human skeletal muscle by a phenornenon called supercompensation (Bergstrom et al. 1967). Carbohydrate supplementation post-exercise increases glycogen resynthesis and this cmresult in glycogen Ievels higher than those present before exercise (Ivy 1991).
In skeletaI muscle al1 glycogenin is attached to the glycogen molecules in a L:l stoichiometric relationship and no reservoirs of free glycogenin exist (Lomako et al.
1990). Lomako, Whelan and coworkers therefore proposed a mechanisrn for supercompensation where the glycogen concentration was increased by transfemng proglycogen into macroglycogen (Alonso et al. 199%). Adamo and Graham (1998) confirmed this hypothesis as they demonstrated that the supercompensation in total glycogen occurred predominantly in the macroglycogen pool but not at the cost of proglycogen. Theoretically, the mode1 of proglycogen as a stable intermediate in the synthesis of macroglycogen suggests that the ratio between proglycogen and rnacroglycogen is predictive of the maximal increase in glycogen levels above basal levels (Hansen et al.
2000). Table 1 shows the calculations of the theoretical increase in glycogen levels dependent on the proglycogen:macroglycogen ratio (Proglycogen is assurned to have a molecular weight of 400kDa and macroglycogen to have a molecular weight of 10,000 kDa). However, in glycogen supercompensated skeletal muscle from both humans and rats a considerable number of the glycogen molecules are still unsaturated (in the proglycogen form), which indicates that the glycogen synthesis stops before al1 proglycogen is converted into macroglycogen (Adarno et al. 1998, Derave et al. 2000,
Hansen et al. 2000).
1.2.7 SYNTHESIS AND GLYCOGENOLYSIS OF PRO- AND MACRO-
GLYCOGEN
To date there are few studies demonstrating the physiological role of pro- and macroglycogen during exercise. Adamo et al. (1998) studied the resynthesis of the two pools of glycogen after exhaustive exercise in humans and showed that proglycogen and macroglycogen differed in both timing and magnitude during resynthesis. Proglycogen was re-synthesized to a greater extent compared to macroglycogen in the early phase of recovery. It was also more sensitive to change in carbohydrate diet compared to macroglycogen. The synthesis of macroglycogen was slower and more constant over a recovery period of 48 hours. When the total glycogen concentration in the muscle reached a concentration over 300-350 rnrnol glucosyl unitdkg dw (supercompensation) the resynthesis of glycogen was exclusively in the macroglycogen pool. These results
demonstrate that proglycogen is not only the precursor of macroglycogen but also the
most dynamic glycogen pool during recovery £kom exercise. These data correlate with the
work by Huang et al. (1997) who used labeled glucose to study glycogen synthesis in
rodent skeletal muscle and were able to demonstrate that the Iabeled glucose first appeared
in the proglycogen pool and then in the macroglycogen pool.
Asp et al. (1999) studied the resynthesis of proglycogen and macroglycogen in hurnan
athIetes afier a marathon race. The major finding fiom this study was that the proglycogen
returned more rapidly back to the pre-race concentration than did the macroglycogen
fkaction. These findings were in agreement with the aforementioned work by Adarno et al. (1998).
As mentioned earlier Lornako and Alonso (Alonso et al. 1995b) proposed a mode1 for glycogen synthesis and degradation where proglycogen was a stable intermediate in the synthesis of mature glycogen (macroglycogen). They speculated that there are two forms of glycogen synthase; proglycogen synthase controlling the synthesis of proglycogen and macroglycogen synthase controlling the synthesis of macroglycogen. In the aforementioned study of the resynthesis of pro- and macroglycogen by Adamo et al.
(1998) the proglycogen pool appeared to be synthesized rapidly when blood glucose and insulin levels were high whereas the synthesis of macroglycogen appeared to be unaffected by the insulin concentrations. In surnmary, the findings fiom this study indicate the existence of two forms of glycogen synthase regulating the synthesis of pro- and rnacroglycogen respectively. Whether proglycogen synthase or macroglycogen synthase are two stnicturally different enzymes or one enzyme under different regdation is at the present time unknown.
Graham et al. (2001) studied the effect of exercise intensity and duration as well as the effect of repeated bouts of exercise on the two pools of glycogen in human skeletal muscle. This study demonstrated that the proglycogen pool is more dynarnic than the macroglycogen pool and that the net rate of glycogenolysis in most exercise situations is greater for the proglycogen pool. When the exercise intensity was increased this caused an increase in the net rate of glycogenolysis of both the pro- and macroglycogen pooI but this increase was more distinct in the proglycogen pool. Repeated bouts of exercise caused a decrease in the net rate of the glycogenolysis with tirne and this was predominately caused by a decreased net rate of catabolism in the macroglycogen pool.
When Asp et al. (1999) studied the utilization of pro- and macroglycogen in skeletal muscle during a marathon race they concluded that a greater fraction of macroglycogen was utilized compared with proglycogen. These findings are in contrast to the work by
Graham et al. (2001). However, the type and intensity of the exercise differs between the two studies. In addition the study perfomed by Asp et al. was a field study with the exercise intensity uncontrolled.
In another study perfomed by Shearer et al. (2001) it was demonstrated that proglycogen and macroglycogen were degraded at different rates depending on the initial glycogen concentration. ProgIycogen was preferentially degraded over macroglycogen at higher initial glycogen concentrations whereas proglycogen and rnacroglycogen contributed equally to the glycogenolysis at lower total glycogen concentrations. This led the authors to specuiate over why proglycogen is preferentially degraded dunng catabolism. The outer branches of the smaller proglycogen molecule may be more readily accessible for glycogen phosphorylase compared to the more densely branched macroglycogen molecule. In addition the number of proglycogen molecules by far outnumber the macroglycogen molecules. Another explanation could be the different subcellular location of pro- and rnacroglycogen within the muscle, resulting in different susceptibility to degradation by glycogen phosphorylase. Friden et al. (1989) demonstrated that glycogen granules were distributed in five distinct subcellular locations and that during exercise the glycogen granules that were preferentially degraded were those located near the Z disks and 1bands.
In contrast Derave et al. (2000) showed that macroglycogen was the major contributor of total glycogenolysis in rats when muscles were glycogen supercompensated. For rodents receiving a nomal or low carbohydrate intake the proglycogen and macroglycogen fiaction contributed almost equally to the glycogenolysis. Furthemore they showed that the initial concentration of macroglycogen but not the proglycogen concentration was significantly corretated to the glycogenolysis.
The results of these experiments demonstrate that both proglycogen and macroglycogen are suitable substrates for glycogen phosphorylase during exercise in both human and rat muscle. The proglycogen pool, however, appears to be the most dynarnic forrn in human skeletal muscle whereas in rodent skeletal muscle the macroglycogen pool is the major contributor of glycogenoIysis at least during high glycogen levels. The hypothesis proposed by Lomako et al. (1995) that proglycogen is a stable intermediate and an arrest point in the glycogenolysis therefore does not seem to be correct. 0-1 -4 linkago krworn nvo glucose uniu
Figure 1. Schematic structure of two outer branches of a glycogen particle with a(l-4) and a(1-6)linkages.
UDP-Glc
"giycogcn initiaor synlhuc" 8nncbi.W- Addùig -8 glucose Rcsiducs to Tyr- 194
Figure 2. Glycogen synthesis on a protein core as proposed by Krisman and Barengo (1975). Glucosyl units (UDP-glucose) are transferred to a protein backbone by a glycogen-initiator synthase to form an oligosaccharide. Further elongation of the glycogen particle is performed by glycogen synthase (GS) and branching enzyme when the oligosaccharide is long enough. Transfii of - 8 giucosyl units dpcdby giycogaiin
Figure 3. The interaction between glycogenin and glycogen synthase in the synthesis of glycogen (Smythe and Cohen 1991). The est step in the biogenesis involves the covalent attachrnent of a glucosyl unit to tyr-194 in glycogenin catdyzed by glycogenin itself (self- glucosylation). Glycogenin and glycogen synthase (GS) fonn a 1:1 complex and the formation of the oligosaccharide primer is catalyzed by glycogenin. When the oligosaccharide primer is 7-11 glucosyl units long fiwther growth of the glycogen molecule is catalyzed by GS and branching enzyme. Macroglycogen (IO7 Da) Proglycogen (400 kDa)
Phosp horylase + Debranching enzyme
4 Glycogen syntase + Branching enzyme I t Gtycogen syntase + Branching enzyme
- 8 glucose residues added bv autocataIvsis Autoglycosyiated giycogenin
Figure 4. Mode1 of glycogen synthesis as proposed by Whelan and colleagues (Alonso et al. 1995b). According to the rnodel glycogenin is auto-glucosylated at Tyr- 194. Proglycogen (400 kDa) is a stable intermediate in the synthesis of glycogen and under norrnal conditions never broken down to glycogenin. Macroglycogen is the hlly glucosylated glycogenin with a rnolecular weight of 10,000 kDa. According to the rnodel two separate glycogen synthase activities are responsible for the synthesis of proglycogen and macroglycogen: proglycogen synthase and macroglycogen synthase. Table 1. Theoretical increase in glycogen levels above basal values- Proglycogen in Macroglycogen in By weight Molar pro:macro Theoretical % of glycogen % of glycogen by pro:macro ratio increase in by weight weight ratio glycogen above basal values O 100 0-00 0.0 1.O 10 90 0.1 1 2.8 3.4 20 80 0.25 6.3 5.8 30 70 0.43 10.7 8.2 40 60 0-67 16.7 10.6 50 50 1-00 25 13 .O 60 40 1.50 37.5 15.4 70 30 2.33 58.3 17.8 80 20 4.00 100.0 20.2 90 10 9.00 225.0 22.6 1.2.8 REFERENCE LIST
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Clzem. 225, 1O3 - 124. CHAPTER 2
The Effect of Extraction Time and Perchloric Acid
Concentration on the Recovery of Pro- and Macroglycogen from Muscle Biopsies fkom the Horse
'Johan T. Brojer, 'Henry R. StWpfli and 'Teny E. Graham
1 Department of Clinical Studies, Ontario Veterinary College, ' Human Biology &
Nutritional Sciences, University of Guelph, Guelph, Ontari NI G 2 W 1, Cananda
For srrbïnission to the Carr adiarz Jorrrït al of Veterhary Researclz 2.0 ABSTRACT
The objectives of this shrdy were to determine whether the concentrations of equine muscle proglycogen (PG) and macroglycogen (MG) are influenced by different extraction times or PCA concentrations. In the first study 20 individual muscle biopsy sarnples were divided into four parts each and then randomly subjected to four different extraction periods (10, 20, 60 and 120 minutes) with 1.5 M PCA. In the second study six individual muscle biopsies were divided into 24 pieces each and then randomly subjected to 12 treatment combinations of different PCA concentrations (0.5, 1.5 and 3.0 M) and extraction times (IO, 20, 30 and 40 minutes). The results frorn study one indicated that
MG and PG concentrations were affected only at the extraction period of 120 minutes; PG decreased (Pc0.05) and MG increased (not significant, PO.05). In the second study extraction in 3.0 M PCA yielded significantly lower PG and higher MG concentrations compared to extraction in 0.5 M md 1.5 M PCA for each of the chosen extraction times
(10-40 minutes). The results fkom this study further support the existence of two glycogen pools and dernonstrate that they are not an extraction artifact. It also suggests that the two pooIs are stable during extraction over a range of acid concentrations (0.5 - 1.5 M) and extraction tirnes. However, if the exposure to acid is very long and/or the acid concentration is high, some of the insoluble PG appears to be hydrolyzed and enters the
MG pool.
2.1 INTRODUCTION
The metabolism of muscle glycogen has been intensely studied in both human and equine exercise physiology. Severd earlier studies conducted on human and rat muscle indicated that glycogen was not a uniform molecule (1-3). Recent studies have shown that
glycogen exists in two molecular fonns, proglycogen (PG) and macroglycogen (MG) (4-
6). The bvo forms have identical protein contents but differ in the arnount of associated carbohydrate. PG is the srndler glycogen molecule with a molecular mass up to 400 kDa whereas MG is the larger glycogen molecule with a maximum molecular weight of approximateiy 10,000 kDa (7). Due to the difference in the ratio of carbohydrate to protein between the two glycogen molecules they cm be separated on the basis of their solubility in acid. Proglycogen was found to precipitate in trichloroacetic acid (TCA) and perchloric acid (PCA) whereas MG was soluble in these acids (6,7). It has also been demonstrated that the relationship between the two pools of glycogen in hurnan skeletal muscle was not influenced by the type of acid (PCA vs TCA) or by the strength of the
PCA in the range between 0.5 and 3 M (3).
It is not clear whether PG and MG exist as two distinct molecular forms or merely represent a continuum of size ranges (4,5,8,9). Despite this, several reports showed that the two foms of glycogen can be separated based on acid solubility in PCA (6,10,11).
Recent studies have indicated that the two glycogen forrns function as different metabolic pools under conditions of glycogenolysis and re-synthesis (1OYl2-l4). These studies indicate that PG and MG are not only differentiated in tems of acid-solubility but also with regard to metabolic regulation.
Recently we have demonstrated that equine muscle glycogen is also composed of MG and PG (1 1). Our results showed that the two foms could be measured in biopsy-sized samples as reliably as total glycogen. The PG and MG were separated by exposing fieeze- dried muscle biopsies to ice-cooled 1.5 M PCA for 20 minutes. The extraction time of 20 minutes has been used boa in our studies of equine muscle as well as in previous studies
of PG and MG in rat (12,15) and human muscle tissue (6,10,13,14,16). However, it is
possible that the two pools could be created as an artifact of the extraction process and the
separation of PG and MG may depend on factors like acid concentration and extraction
tirne. No information is currently available in the literature concerning stability of the two
foms of glycogen in relation to extraction time. If the relationship between recovered pro-
and rnacroglycogen continues to change with extraction tirne without reaching a stable
level, the reliability of the separation technique rnust be questioned and the biological
significance of the data would be limited. Information about the effect of the combination
of different extraction times and PCA concentrations on the relationship of PG and MG is
also lacking, Before studies of the physiological response of glycogenolysis, synthesis and
regulation of the two glycogen foms can be conducted in the area of equine exercise
physiology the above rnentioned issues have to be fiirther addressed.
The purpose of this study was to evaluate the influence of extraction time and PCA
concentration on the amount of PG and MG recovered frorn fieeze-dried muscle biopsy samples fiom horses. We hypothesized that the PG and MG pool would rernain constant over a wide range of PCA concentrations and extraction times.
2.2 MATERIALS AND METHODS
Horses
Ten Standardbred horses (8 mares and 2 geldings, age 2-10 years) were used in study
1 whereas six O ther Standardbred horses (5 mares and 1 gelding, age 2-1 1 years) were used in study 2. Al1 horses were clinically healthy. The care and use of anirnals followed the Guide to Care and Use of Experimental Animals (Canadian Council on Animal Care,
Ottawa, Ontario). Al1 animal experiments were conducted after approval by the Animal
Care Cornmittee of the uni ver si^ of Guelph and perfïormed in cornpliance with their
recomrnendations.
Ex~enmentaldesign of studv 1 - Effect of extraction time
The effect of different extraction tirnes on the recovery of MG and PG were evaluated
for the MG+PG determination technique. Ten individual fieeze dried and powdered
muscle biopsies were divided into four equal portions each, and were randomized into
four different treatment combinations. To assess influence of extraction time the four parts
were extracted in 1.5 M PCA as in the original method (6)for detemination of MG and
PG but for 10 min, 20 min, 60 min and 120 min respectively. Extraction tirne was measured fiom the time PCA was added until the commencement of centrifiigation.
Experimental design of study 2 - Effect of the combination of different PCA concentrations and extraction times
The influence of different PCA concentrations and extraction times on the recovery of
PG and MG fiom fieeze dned muscle biopsy sarnples were studied. Six individual freeze- dried powdered muscle sarnples were divided into 24 pieces each. These pieces of each sarnple were randornly subjected to 12 different treatrnent combinations of varying PCA molarity (0,5 M, 1.5 My and 3.0 M) and extraction time (10, 20, 30 and 40 minutes).
Extraction tirne was measwed fiom the tirne PCA was added until the commencement of centrifugation. Muscle biopsies
Muscle biopsy sarnples were obtauled by needle biopsy fi-om the middle gluteal muscIe as described by LindhoIm and PiehI (17). The biopsy site was 15 cm from the tuber coxae on a line to the base of the tail and at a depth of 6 cm. On each occasion two sarnples were obtained from one biopsy site. The same skin incision was used for both biopsies. Al1 sarnples were frozen ùnmediately in liquid nitrogen and stored at -80°C until analyzed. The sampIes were fieeze dried, dissected free from blood, connective tissue and fat. The two sarnples fkom each horse were pooled together and powdered before analysis.
Analysis
Proglycogen and MG fractions were separated with the method described by Adarno and Graham (6) with modification of the extraction times and PCA concentrations.
Bnefly, in Adarno and Graham's method 200 pl of ice-cooled PCA was added to 2.0 - 2.5 mg freeze dried muscle sarnples. The muscle samples were rnixed with PCA using a plastic rod. The extraction was performed on ice followed by centrifugation at 3000 revolutions/rnin for 15 minutes at 4OC. One hundred pl of the supernatant was kept for the detemination of MG and the peIlet used for the determination of PG. The fractions of PG and MG were boiled in 1 M HCI for 2 hours and the formed glucosyl units were subsequently measured fluorometrically with the hexokinase method (18). Values are reported as glucosyl units per kg dry weight (dw).
In study 2 an aliquot was taken fiom each of the PCA extracts (MG fractions) before hydrolysis of the MG and PG &actions in HCI. The aliquots were neutralized with KHCO, (19) and the samples used for determination of free glucose using an enzymatic
fluorometric rnethod (1 8).
Statistical analysis and calculations
Al1 data were subjected to analysis of variance using the General Linear Mode1
procedure in the computer software Minitab for Windows (0 1997. Minitab Inc. State
College, PA, USA). The following model was used for study 1: Y, = p + ri + p, + E~, where Y, is the observation, p is the mean value, ri is the effect of extraction time, pj is the effect of horse (block effect) and E,, is the experirnental error. In study 2 the following model was used: Y,,, = p + pi + aj+ Bk + (aB)j, + cik+ Ôiixi, where Yi*,is the observation, p is the mean value, pi is the block effect (horse), ajis the effect of extraction tirne, Pk is the effect of PCA concentration, (np),, is the effect of interaction between extraction time and PCA concentration, E,, is the experimental error and 6,,, is the sampling error. The significance level was set at P<0.05. The differences between each of the treatments where compared by Tukey test. Values are presented as means + SE. Total glycogen (GJ was obtained by adding the two fractions of glycogen (G, = PG + MG).
2.3 RESULTS
Study 1 - Effect of extraction time
The results from this study indicate that extraction time had little effect on the concentration of MG and PG obtained fiom keeze dried muscle samples extracted in 1.5
M PCA. The PG and MG concentrations were not affected by extraction tirnes between 10 and 60 minutes (Table 1). Extraction during 120 minutes showed a tendency towards decreased PG and increased MG concentration. The average decrease in PG was 37,
which is very similar to the average increase in MG of 17. The decrease in PG was
statistically significant but the increase in MG was not. The total glycogen values were
very sirnilar between the different extraction times and were not statistically significant.
Study 2 - Effect of the combination of different PCA concentrations and extraction times
Similar values for PG and MG were obtained with extraction in PCA concentrations of 0.5 M and 1.5 M at each extraction tirne (10-40 minutes) (Table 2 and 3). Extraction tirne had a significant effect on PG and MG concentrations at the level of 40 minutes when muscle biopsies were extracted in 3.0 M PCA. Extraction in 3.0 M PCA yielded significantly lower PG concentrations and higher MG concentrations (compared to 0.5 M and 1.5 M) for each extraction time.
Total glycogen concentrations were sirnilar over the range of PCA concentrations and extraction times with one exception; extraction during 20 minutes yielded modestly but significantly higher G, vaIues for PCA concentration of 3.0 M compared to 0.5M and
1.5M (Table 4). Analysis of glucose in al1 aliquots trom the MG Eactions before hydrolysis in HC1 yielded low values (0.14 - 0.89 molglucose per kg dry weight). The mean glucose concentration in each of the 12 treatment combinations of extraction time and PCA concentration were very similar and ranged behveen 0.45 and 0.54 mm01 gIucose per kg dry weight. In conclusion the glycogen was not hydrolysed into glucose residues during the PCA extraction procedure for any of the PCA concentrations or extraction times. 2.4 DISCUSSION
The purpose of this study was to determine whether the two structural forms of muscle glycogen are stable during the extraction procedure with regard to extraction time and PCA concentration. Our data indicate that extraction time has little effect on the concentration of MG and PG obtained Erom fieeze-dried muscle samples. There was no detectable time effect on PG or MG extracted in 0.5 M or 1.5 M PCA in study 2.
However, the maximum extraction tirne was limited to 40 minutes. The concentrations of
PG and MG were also stable when extracted in 3.0 M PCA for up to 30 minutes- It appears that very long extraction times are necessary to affect the relationship between the two pools of glycogen in muscle tissue when extraction is performed in 1.5 M PCA. In. study 1 the extraction time was prolonged to 120 minutes. After 120 minutes of extraction there was a small decrease in PG and a similar increase in MG, but the change in concentration was only significant for PG. Considering that extraction of muscle metabolites in 0.5 M PCA is usually performed during 10 to 20 minutes (19), an extraction time of 120 minutes must be considered to be extremely long and probably does not have any physiological relevance. Our data dernonstrate that under normal circumstances there is no evidence that the extraction technique generates artifacts that change the concentrations of the two pools and we can therefore attach biological significance to them.
We were not able to identiQ any differences in PG and MG between extraction in 0.5
M and 1.5 M PCA. Extraction in 3.0 M PCA yielded significantly lower PG and higher
MG concentrations for al1 extraction times (between 10 and 40 minutes). It may be that the stronger acid or longer exposure caused a modest amount of hydrolysis of small chains of polysaccharides from the insoluble PG fraction and they entered the MG kaction. This
appears to be the case as the change in each pool tends to be inversely related. These results differ from the data presented by Jansson (3) who showed that the reIationship between PCA soluble and -insoluble glycogen was not influenced by the PCA in the range between 0.5 and 3 M. It is possible that species variation accounted for the differences found; the study by Jansson was performed on hurnan skeletal muscle tissue whereas our study was conducted on equine skeletal muscle. In addition, Jansson's study was performed only with an extraction time of 20 minutes. Extraction in 3 M PCA during longer extraction times could have resulted in elevated MG and lower PG concentrations.
Analysis of glucose in the PCA extracts (MG kactions) before acid hydrolysis in HCl in the second study showed that none of the glycogen fiactions were hydrolyzed into glucose residues by the PCA regardless of extraction times and PCA concentrations.
These results exclude the possibility of pre-hydrolysis of glycogen into glucose residues by the 3.0 M PCA as an explanation for the differences in the relationship between PG and MG obtained by this PCA concentration. It is possible that the higher PCA concentration (3 M) extracted more MG from the samples and that the extraction in 0.5 M and 1.5 M PCA was incomplete. This is however not likely. Extraction of muscle metabolites fkom fieeze-dned muscle is considered to be cornplete after extraction for 10 to 20 minutes in 0.5 M (19). If the extraction had been incomplete one would not expect such a stable relationship between PG and MG as that obtained with different extraction times. The relationship between the two forms of glycogen was also very stable in both
0.5 M and 1.5 M PCA. The intracellular concentration of glucose and glucose-6-phosphate (G6P) are low in muscle tissue during rest (20). Under conditions of intense exercise the levels of intracellular glucose and G6P rises and since these metabolites will be recovered in the
MG fraction dunng the extraction procedure they could cause a significant overestimation of the MG concentration. When the glucose residues formed after HC1 hydrolysis were measured in the MG fiaction by the hexokinase method the G6P was excluded from the
MG fiaction by incorporating the enzyme GoP-dehydrogenase in the reagent and not adding the enzyme hexokinase until after the initial reading of NADPH. Separate analysis of intracelluIar glucose in post exercise sarnples is however required and enables exclusion of this error by subtracting glucose from the MG concentration.
There is controversial idormation in the literature on whether PG MG exist as a distinct molecular entity or if the two pools in fact are a continuum of molecular sizes in the range from glycogenin up to a large MG molecule of approximately 10.000 kDa
(4,5,8,9). Irrespective of the range of rnolecular sizes of PG and MG several studies have demonstrated that the two fkactions cm be separated based on acid solubility (6,10,11).
Recent studies have indicated that the two glycogen foms function as different metabolic pools under glycogenolysis and re-synthesis (1 0,12-14). Furthemore, the wide variation found in the proportion of PG to MG between different tissues indicates that there are factors that control the relative arnounts of PG and MG (5). The fact that the PG and MG pools are very stable during extraction in variable acid concentrations and extraction times further supports the theory of the existence of two molecular forms of glycogen that can be reliably separated. The separation technique for PG and MG allows Mer understanding of the
metabolism of muscle glycogen in the field of exercise physiology. The technique also
permits determination of total glycogen, PG, MG and metabolites kom the sarne piece of
tissue, which eliminates the variation between muscle pieces. It is also possible that this
technique could be of value in the investigation of equine glycogen storage myopathies.
In surnmary, the present study demonstrated that the separation technique for PG and
MG is reliable and not simply an artifact generated by extraction time andor acid concentration. The concentration of the two glycogen pools were very stabIe with extraction time and concentration of PCA in the range of 0.5 and 1.5 M. Extraction in 3.0
M PCA yielded significantly different values for PG and MG and this acid concentration is therefore not considered to be useful for separation of the two glycogen pools in equine skeletal muscle.
2.5 ACKNOWLEDGEhlENTS
EP Taylor Equine Trust Fund generously supported this study. We would like to thank Kim Kultiman, Swedish Veterinary Institute for advice with the statistical analyses. Table 1: Mean pro-, macro- and total glycogen concentrations for different extraction times.
Extraction tirne
Glycogen analyses 10 min 20 min 60 min 120 min
Gt 530.9 $: 12.4 537.6 + 23.4 516.3 t 11.3 510.5 + 5.7 Values are mean + SE, n=10. Al1 values are in molglucosyl unitskg dry weight, PG is proglycogen, MG is macroglycogen and Gr is total glycogen. G, was obtained by adding the two Eiactions, PG and MG- * Significantly different (P<0.05) fi-orn extraction time 10 and 20 min. Table 2: Mean proglycogen concentration for different combinations of extraction tirne and perchloric acid (PCA) concentration.
Extraction time PCA 0.5 M PCA 1.5 M PCA 3.0 M 10 min 327.2 f 11.4 325.6 + 11.8 307.3 t 11.2* 20 min 323.7 t 10.9 327.6. + 7.7 304.7 + 12.1* 30 min 319.3 + 11.2 322.3 I9.5 294.4 + 11.2* 40 min 313.4 t 11.7 313-4 t 10.4 281.6 I12,8* ' Values are mean t SE. Each data set contains values fiom 6 horses with 2 sarnpling analyses per horse (n=12). Al1 values are in mm01 glucosyl unitskg dry weight. *Significantly different (P c 0.05) korn PCA concentrations 0.5 M and 1.5 M at each level of extraction time. " Significantly different (P < 0.05) frorn extraction time 10 min and 20 min at the level of PCA concentration 3.0 M. Table 3: Mean macroglycogen concentration for different combinations of extraction time and perchlonc acid (PCA) concentration.
Extraction time PCA 0.5 M PCA 1.5 M PCA 3.0 M 10 min 138.0 + 18.2 141.8 + 19.8 172.3 + 26-4* 20 min 138.9 + 18.3 146.7 t 20.0 184.9 + 27.2" 30 min 142.9 rt 19.2 152.1 4 21.5 184.8 i 26.G* 40 min 142.4 i 18.8 149.8 Ir 21-7 190.2 I 26.5* * Values are mean f: SE. Each data set contains values f?om 6 horses with 2 sampling analyses per horse (n=I 2). Al1 vaIues are in mm01 glucosyl unitdkg dry weight. *Significantly different (P < 0.05) fiom PCA concentrations 0.5 M and 1.5 M at each Ievel of extraction tirne. ' Significantly different (P < 0.05) fiom extraction time 10 min at the level of PCA concentration 3 .O M. Table 4: Mean total glycogen concentration for different combinations of extraction time and perchloric acid (PCA) concentration.
Extraction time PCA 0.5 M PCA 1.5 M PCA 3.0 M 1O min 465.2 + 22.5 467.4 I 2 1.7 479.6 + 27.8 20 min 462.6 + 18.3 474.3 + 21-5 489.6 t 28.2* 30 min 462.3 t 24.9 474.4 t 24.3 479.2 + 25.6 40 min 455.8 t 26.0 463.2 + 24.9 47 1.9 t 26.2 Values are mean I SE. Each data set contains values from 6 horses with 2 sarnpling analyses per horse (n=12). Al1 values are in mm01 glucosyi unitskg dry weight. *Significantly different (P c 0.05) Eom PCA concentrations 0.5 M at the level of 20 min extraction time. 2.6 REFERENCES
1. Kits Van Heijningen AJM, Kemp A. Free and fixed glycogen in rat muscle.
Biochem J l955;59:487-491.
2. Stetten Jr. D, Stetten MR. Glycogen Metabolism. Physiol Rev 1960;40:505-537.
3. Jansson E. Acid soluble and insoluble glycogen in human skeletal muscle. Acta
Physiol Scand 1981;113:337-340.
4. Alonso MD, Lomako J, Lomako WM, Whelan WJ. A new look at the biogenesis of
glycogen. FASEB J 1995;9:1126-1137.
5. Lomako J, Lomako WM, Whelan WJ, Dombro RS, Neary JT, Norenberg MD.
Glycogen synthesis in the astrocyte: f?om glycogenin to proglycogen to glycogen.
FASEB J 1993;7:1386-1393.
6. Adamo KB, Graham TE. Comparison of traditional measurements with
macroglycogen and proglycogen analysis of muscIe glycogen. J Appl Physiol
7. Lomako J, Lomako WM, Whelan WJ. Proglycogen: a low-molecular-weight fom of
muscle glycogen. FEBS Lett 199 1;279:223-228.
8. Lomako J, Lomako WM, Whelan WJ. Glycogen metabolism in quail embryo
muscle. The role of the glycogenin primer and the intermediate proglycogen. Eur J
Biochem 1995;234:343-349. 9. Roach PJ, Skurat AV. Self-glucosylating initiator proteins and their role in glycogen
biosynthesis. Prog Nucleic Acid Res Mol Bi01 1997;57:289-3 16.
10. Adarno KB, Tarnopolsky MA, Graham TE. Dietary carbohydrate and postexercise
synthesis of proglycogen and macroglycogen in human skeletal muscle. Am J
Physiol 1998;275:E229-334.
11. Brojer, JT. Proglycogen and macroglycogen in equine skeletal muscle FISCThesis].
Guelph, Ontario: University of Guelph, 2001.
12. Derave W, Gao S, Richter EA. Pro- and macroglycogenolysis in contracting rat
skeletal muscle. Acta Physiol Scand 2000;169:29 1-296.
13. Graham TE, Adamo KB, Shearer J, Marchand 1, Saltin B. Pro- and
rnacroglycogenolysis: relationship with exercise intensity and duration. J Appl
Physio12001;90:873-879.
14. Shearer J, Marchand 1, Tarnopolsky MA, Dyck DJ, Graham TE. Pro- and
rnacroglycogenolysis during repeated exercise: roles of glycogen content and
phosphorylase activation. .iAppl Physiol2001;90:880-888.
15. Hansen BF, Derave W, Jensen P, Richter EA, No limiting role for glycogenin in
determining maximal attainable glycogen levels in rat skeletal muscle. Am J Physiol
Endocrinol Metab 2000;278:E398-404. Asp S, Daugaard JR, Rohde T, Adarno K, Graham TE. Muscle glycogen accumulation after a marathon: roles of fiber type and pro- and macroglycogen. J
Appl Physiol 1999;86:474-478.
Lindholm A, Piehl K. Fibre composition, enzyme activity and concentrations of metabolites and eIectrolytes in muscles of standardbred horses. Acta Vet Scand
1974; 1S:287-309.
Bergmeyer, H. Methods of Enzymatic Analysis. New York: Academic,
1974;3:1128-1131
Harris RC, Hultman E, Nordesjo LO. Glycogen, glycolflic intermediates and high- energy phosphates determined in biopsy sarnples of muscuIus quadriceps femoris of man at rest. Methods and variance of values. Scand J Clin Lab Invest 1974;33: 109-
120.
Nevill ME, Boobis LH, Brooks S, Williams C. Effect of training on muscle metabolism during treadrnill s~rintin~.J ADDI Phvsiol 1989:676:2376-2382. CHAPTER 3
Analysis of Pro- and Macroglycogen in Muscle Biopsies
fkom Horses
'J T Brojer, 'H R Stampfli and 'T E Graham
' Department of Clinical Studies, Ontario Vetennary College and ' Human Biology and
Nutritional Sciences, University of Guelph, Guelph, Ontaria, Cananda
Fur srr btriissiotr ru the rlnteriuuz JOrrrizul of Vetevirzary Researclz 3.0 ABSTRACT
Objective - To establish whether proglycogen (PG) and macroglycogen (MG) exist in equine skeletal muscle and to compare two different analytical methods (acid hydrolysis,
AC, and PG+MG determination) for measurement of total muscle glycogen (Gly,J Eom biopsy sarnples.
Animals - Clinically normal horses of different breeds.
Procedure - In the first study 45 individual muscle biopsies f?om the middle gluteal
(n=31) or triceps muscle (n=14) were analyzed using both AC and MWPG determinations for Gly,,,, The variability within muscle biopsies for the AC and MGtPG methods were calculated fkom duplicate analyses of the muscle sarnples. In the second study the variation in MG and PG between muscle biopsies and the effect of sampling depth on the concentration of MG and PG in the middle gluteal muscIe was studied.
Results - There was a strong correlation between Gly,, values obtained by AC and
MGRG determination (r=0.99). Coefficients of variation for within and between biopsy variability of Gly,,, were approximately 4% for both methods. The PG fraction was always in excess of MG. However, with increases in Gly,,, the percentage of MG increased as well. The superficial biopsies froom the middle gluteal muscle contained significantly
(P<0.05) more Gly,,, and PG than the deeper biopsies.
Conchsions and clinical relevance - These studies confirm that MG and PG exist in equine skeletal muscle and that they can be measured reliably in biopsy-sized samples.
This new technique could be applied in future studies to investigate the glycogen metabolism in horses during exercise. 3.1 INTRODUCTION
The metabolism of muscle glycogen has been intensely studied in both human and
equine exercise physiology. The introduction of the percutaneous muscle biopsy
technique b y Bergs troml has enabled Our understanding of both equine and human muscle physiology to expand tremendously over the last 30-40 years.
Earlier biochemica12" and electron rnicroscopic5 studies indicated that glycogen was not a uniforrn molecule. Recent sîudies have identified two pools of glycogen in human and rat skeletal muscle: proglycogen (PG) and macroglycogen (MG)." Proglycogen is the smaller molecule with an estimated molecular weight of up to 400 kDa, whereas MG is the larger mature glycogen molecule with a maximum molecular weight of approximately
10 000 k~a.~They have similar protein content of approximately 40 kDa but different amount of associated carbohydrate. Thus, on a weight basis PG is made up of approximately 10% protein whereas the MG contains only 0.4% protein. As the two forms differ in their protein to carbohydrate ratios, they cm be separated based on their acid solubility in perchlonc acid (PCA) or trichloroacetic acid (TCA). Macroglycogen is soluble in PCA and TCA but PG is insoluble and precipitates in these acids due to its higher protein content.' Recently, Adamo and Graham8 validated a separation technique for MG and PG for human and rat skeletal muscle in which fteeze-dried muscle biopsies were exposed to 1.5 M PCA at a temperature of 0°C.
Human resting skeletal muscle contains approximately 6575% of the total glycogen in the PG f~rrn'.~~whereas resting rodent muscle contains 85-90% PG.'." Under most exercise conditions the net rate of glycogenolysis is higher for the PG pool compared to the MG pool in human skeletal muscle.'0*" In contrast, Derave et al." have demonstrated that in rodent skeletal muscle MG is degraded to a greater extent compared to PG during glycogenolysis. These findings indicate that there are species differences with regard to both metabolism and relationship between PG and MG in skeletal muscle. Proglycogen and macroglycogen have not been examined in equine muscle. The separation technique therefore requires Further validation for equine skeletal muscle before studies of the physiological response of glycogenolysis, synthesis and regulation of the two forms of glycogen can be conducted in exercising horses.
Earlier studies in both human and rat muscle have dernonstrated that the relative proportions of MG and PG are dependent on the total glycogen on cent ration.^^"^" With increasing total glycogen concentrations the relative proportion of MG increases.
Glycogen is found in higher quantities in equine skeletal muscle compared to rat and hurnan muscle. We hypothesized that equine skeletal muscle is composed of PG and MG and that the relative proportion of MG is higher in equine muscle compared to rat and human muscle.
The aims of this study were 1) to establish if MG and PG exist in horse muscle; 2) to compare analysis of total glycogen by AC and MWGdetermination; 3) to evaluate the precision of the AC and MGtPG determination technique and 4) to evaluate the effect of sarnpling depth in the glrtteus medius muscle on the concentration of MG and PG.
3.2 MATERLkLS AND METHODS
The care and use of animals followed the Guide to Care and Use of Experimental
Animnls (Canadian Council on Animal Care, Ottawa, Ontario). Al1 animal experiments were conducted after approval by the Animal Care Cornmittee of the University of Guelph
and performed in comphnce with their recommendations.
Expenment 1 - Within biopsy variability and cornparison between AC and MG+PG detenninations
A total of 45 individual muscle biopsies were used in this study. The biopsies were collected f?om the gluteus medius muscle or the triceps muscle during conditions ranging from rest to 12 hours post-exercise in order to obtain biopsies with a wide variation in glycogen content. The glutezrs medizcs muscle was biopsied at rest fkom 24 clinically normal Standardbred horses (20 mares and 4 geidings, age 2-17 years) and fiom 7 crossbred Arabian horses (1 mare and 6 geldings, age 8-17 years) approximately 12 hours after completion of an endurance ride. Muscle biopsies fiom the triceps muscle were obtained from 14 clinically normal Standardbred horses at rest (1 1 mares and 3 geldings, age 2 -11 years). Total glycogen was measured in the muscle biopsies (n=45) using two different methods, AC and MG+PG determinations. The error of the analytical method
(variability within muscle biopsies) for the AC and the MG+PG method was calculated fiom duplicate analyses of the individual muscle samples (n=45).
Experiment 2 - The effect of sarnpling depth in the ghtezrs meditrs muscle and the between biopsy variability for PG and MG
Nine Standardbred mares (age 6 -15 years) were biopsied at rest from the nght and lefi gluteus medius muscle. Two repeated biopsies were obtained on the same occasion at a depth of 6 cm fiom the right glureus medizts muscle for evaluation of between biopsy variability. Two biopsies were collected fiom the lef? gluteus medizts muscle; one at a depth of 40 mm and one at a depth of 80 mm. The first biopsy was taken at the 40 mm depth. Glycogen was analyzed using the MWPG determination.
Muscle biopsies:
Muscle biopsy sarnples were collected fiom the glutezts mediru muscle at a depth of 6 cm, unless othenvise stated, and from the triceps muscle at a depth of 4 cm, according to the technique of Lindholm and ~ieh1.l'The biopsy site for the ghteza medius muscle was
15 cm caudomedial to the tuber coxae on a line extending fiom the caudal aspect of the ruber coxue to the base of the tail. The biopsy site for the triceps muscle was located by drawing a 10 cm long perpendicular line nom a point one third of the distance Eom the most caudal part of olecranon to the trtberntZztm majus, pars craniulis of the hztnzerus.
When repeated biopsies were taken the same skin incision was used for both biopsies.
Al1 sarnples were fiozen irnrnediately in liquid nitrogen and stored at -80°C. The samples were fieeze dried, dissected fiee kom blood, connective tissue, and fat, and then powdered before analyses.
Analysis
The AC method for determination of glycogen was adopted fiom Passonneau and
Lauderdale.16 Freeze-dried muscle sarnples weighrng between 1.5 and 2 mg were boiled in
1 M HC1 for 2 hours foIlowed by neutralization with 1 M NaOH. The samples were vortexed and centrifuged at 3000 revolutions/min for 5 minutes. The supernatants were used for fluorometric determination of formed glucosyl units." Proglycogen and MG fiactions were separated based on solubility in PCA with the
method described by Adamo and Graham.' Briefly, 200 pl of ice-cooled 1.5 M PCA was
added to 1.5-3 mg fieeze-dried muscle sarnples. The muscle sarnples were rnixed with
PCA using a plastic rod- The extraction was per£ormed on ice over 20 minutes followed by centnfùgation at 3000 revolution/rnin for 15 min. One hundred pl of the supernatant was kept for the detennination of MG. The remaining PCA was discarded and the pellet used for determination of PG- The PG and MG samples were boiled for 2 hours in I M
HCI and subsequently neutralized with 2 M Trima base, vortexed and centrifûged for 5 min at 3000 revolutions/min. Subsequently, the glucosyl units formed in the supernatants
Eom each fiaction were measured as described above for glycogen.
Calculations and Statistical Analysis
Total glycogen for the MG+PG detennination was calculated as sum of measured PG and MG. The fraction of MG was obtained by dividing the concentration of MG by total glycogen (MG/NG+PG]) and reporting it as a percent (x 200%). Results were analyzed using the cornputer sohvare Minitab for Windows." Linear regression analysis was used to deterrnine correlation among methods for total glycogen determination. Agreement between analyses methods and assessrnent of repeatability (coefficient of repeatability,
CR) were calculated as described by Bland and ~1trnan.I'The lirnits of agreement were calculated as mean difference * 2 standard deviations (ZI~SD). The CR values were obtained by taking twice the standard deviation of the differences of the duplicates. The standard deviation for duplicate analysis was calculated according to the formula SD = JEand the coefficient of variation (CV) was calculated as SD/X where d is
the difference between duplicate analyses, n is the nurnber of duplicates and X is the
mean value. Paired Student's t-test was used to compare mean values of AC, MG+PG,
MG and PG rneasurements between repeated biopsies. Al1 results are expressed as mean
I: SD. The nul1 hypothesis was rejected at P<0.05.
Expenment 1 - Within biopsy variability and cornparison between AC and MG+PG
determinations
This study presents evidence that the MWPG determination is comparable to the AC
method of analysis of total glycogen content in muscle biopsy sarnples fkom the horse
(Fig 1). There was an excellent correlation between the MG+PG and the AC determinations (r=0.99). The line of identity with a dope equal to one and with the intercept of O was almost identical to the regression line (y=x-9.2). The rnean difference for the analyses of total glycogen between the MGPG and AC determination was -7.1 mm01 glucosyl units per kg dw with 95% confidence interval -1 1.2 to -3.0 (Fig 2). The lirnit of agreement was -34.2 to +20.2 mm01 glucosyl units per kg dw.
The relationship between MG and total glycogen (MGtPG) appeared to be curvilinear
(Fig 3). With an increase in total glycogen concentration the percentage of MG increased, but the MG concentration never exceeded the PG concentration. In the muscle biopsy with the lowest total glycogen content (138.8 mol glucosyl unitskg dw) the MG concentration accounted for 19% of Gly,, whereas MG accounted for 43% of Gly,, in the muscle biopsy with the highest total glycogen concentration (578.6 mm01 glucosyl
units/k,o dw)-
The precision, reported as SD, CV and CR, of total glycogen (within muscle biopsy
variability) determined with the AC or MWPG technique was very similar both between
methods and between different concentrations of Gly,,, (Table 1).
Experirnent 2 - The effect of sampling depth in the ghtezrs medius muscle and the
between biopsy variability for PG and MG
The between biopsy variability, reported as SD, CV and CR, for MG, PG and
MWPG was slightly higher compared to the values for the within muscle biopsy
variability (Table 2). The average fiaction of MG to Gly,, was 41%. There were no
significant differences in glycogen content between the repeated biopsies for the
determination of MGtPG, MG and PG (P<0.05).
The sampling depth in the glztteza rnenizrs muscle had a significant effect (P<0-05) on
the concentration of Gly,, and PG (Table 3). Biopsies obtained more superficially had
higher total glycogen and PG content compared to the deeper biopsies. The mean MG
content was also higher in the superficial biopsies, but these differences were not
statistically si,pificant (PcO.05). The average fiaction of MG to Gly,, at the two sampling
depths was almost identical (39.4% and 39.9% respectively).
3.4 DISCUSSION
The main purpose of this study was to demonstrate whether MG and PG exist in horse skeletal muscle and to compare analysis of total glycogen deterrnined by the AC and MG+PG methods over a wide range of glycogen concentrations. The precision of the
analyses of total glycogen, PG and MG fiom muscle biopsy sarnples was also evaluated.
To our knowledge, this study presents the first results for MG and PG in equine skeletal
muscle tissue. Our data strongly indicates that there are two forms of glycogen in equine
muscle and that they can be reliably measured in biopsy-sized sarnples. The relative
proportion of MG to Gly,, is higher in equine muscle (- 40%) compared to rat (10-15%)
and human (25-35%) muscle.
The first study showed an excellent correlation between the AC and MG+-PG methods
for determination of total glycogen over a wïde range of glycogen concentrations. This
result is in agreement with the data for rat and human muscle presented by Adamo and
Graham.* However, the use of correlation does not give sufficient information about
agreement between two different measurement techniques. We therefore used an alternative statistical approach for assessing agreement between methods." This statistical analysis can also be used for assessrnent of reproducibility (coefficient of repeatability).
The differences between the AC and MGRG method did not Vary in any systematic way over the range of measurements (Fig 2). The lack of agreement between the two methods was -34.2 to 20.2 rnrnol glucosyl units per kg dw within the Gly,,, concentration range of 138.8-593.6 molglucosyl units per kg dw. Thus, the MG+PG methodology may give readings for glycogen that are 34.2 molglucosyl units per kg dw beIow or
20.2 rnrnol glucosyl units per kg dw above the AC method in 95% of the measurements.
These values are of the same order of magnitude as the limits of agreement (CR) for duplicate analyses of total glycogen content analyzed by AC or MG+PG (Table 1). Our results of reproducibility (SD, CV) for determination of total glycogen by AC or MG-WG were in good agreement with previous s~xdies.~~~~~~~-~~In addition, the methodological error of the analysis of PG and MG was low and it did not Vary over the range of total glycogen concentrations. We therefore conclude that the MGtPG technique is a reliable method for analysis of total glycogen in biopsy sarnples and that the two methods are comparable.
Previous studies conducted on hurnan and rat muscle have demonstrated that the relative proportions of PG and MG on a weight basis were dependent on total glycogen concentrati~n.~*~'.'~As total glycogen concentration increased in the muscle the proportion of MG increased whereas the proportion of PG decreased. Despite a decrease in the relative proportions of glycogen, the absolute PG concentration increased as total glycogen increased. A similar relationship has been demonstrated in this study for equine skeletal muscle (Fig 3). However, in hurnan skeletal muscle the PG concentration appears to reach its maximal concentration (250-300 mm01 gIucosyl units per kg dw) at total glycogen concentrations of approximately 300-350 mm01 glucosyl units per kg dw.I4
Thus, for human skeletal muscle the increase in total glycogen concentration over 300-350 mm01 glucosyl units per kg dw was predominantely due to increase in the MG pool. This pattern was not identified for equine skeletal muscle in this study as the PG concentration continued to increase up to 350 mm01 glucosyl units per kg dw without reaching a plateau
(data not shown). It is possible that PG reaches a plateau at higher total glycogen concentrations (>600 gIucosyl units per kg dw) but to study this hypothesis muscle biopsies with higher total glycogen concentration than those available in this study are required.
Horses have approximately 40% of their muscle glycogen concentration in the MG form. Assuming a molecular mass of 400 kDa for proglycogen and 10 000 kDa for macroglycogen, the macroglycogen rnolecule will be approximately 25 times the size of a
proglycogen molecule.' Therefore, when the glycogen forms are compared in molar
concentrations approximately 2.7% of the glycogen molecules in the equine gluteus
medius muscle are in the MG form and the proglycogen molecules by far outnumber the
macroglycogen molecules. This could have important implications for the utilization of
glycogen during exercise. The PG pool has been shown to be the rnost dynamic pool
during exercise in humans and it is associated with a higher rate of glycogenolysis in most
situations of exer~ise.'~Macroglycogen is a very densely branched molecule and it has been demonstrated that large glycogen molecules contain regions that are resistant to degradation with a-arnyla~e.~'It is therefore possible that MG is more resistant to glycogen phosphorylase and debranching enzyme cornpared to the smaller much more abundant PG rnolecufe.
There is controversial information in the literatwe regarding whether PG and MG exist as discrete rnolecular forms or as a range of ~izes.~.'"*'-' However, the interpretation of our results is not dependent on this issue, Our data show that the two fiactions of glycogen found in horse muscle can be separated based on acid solubiIity, regardless of the molecular sizes. This is in agreement with previous work in human and rat rnus~le.~~''
Jansson4 demonstrated that the relationship between PCA-soluble and -insoluble glycogen was not influenced by the strength of the PCA (range between 0.5 to 3 M), the type of acid (PCA vs. TCA), the keeze-mg procedure or the weight of the samples (0.2-2 mg).
In addition, we have recently demonstrated that the two pools of glycogen are stable during wide variations in extraction times (10-40 minutes) (unpublished observations
Brojer et al.). Thus, the separation of PG and MG by acid solubility is not an extraction artifact caused by factors like the strength of the acid andlor extraction time and we can therefore attach biological significance to thern.
Endogenous tissue glucose and glucose-6-phosphate (G6P) are recovered together with MG in the supernatant during the PCA extraction procedure and they will therefore incorrectly be measured as MG. In resting muscle the concentration of these metabolites are very small relative to the MG fraction. However, during intense exercise the intracellular concentration of glucose and G6P increases,'" which could potentially create a sipificant error in the measured concentration of MG. It is therefore important that, for post-exercise sarnples, endogenous tissue glucose and GoP are rneasured in the PCA supernatant before HC1-hydrolysis and subsequently subtracted from the MG fraction.
When the glucosyl units were measured fluorometrïcally in our study, the enzyme G6P- dehydrogenase was incorporated in the reagent and the enzyme hexokinase was not added until after the initial reading.17 Thus, the measured MG concentration in our study contained intracellular glucose but not intracellular G6P. The post-exercise sarnples in Our study were obtained several hours after the endurance ride was cornpleted, which would give negligible values of intracellular glucose.
The results fi-orn experirnent 2 showed lower total glycogen concentrations in the deeper compared to the superficial biopsies from the ghteus rnenitrs muscle. This is most likely a result of the non-uniform fiber composition within the glutais medius muscle; the highest proportion of type LI fibers is found superficially, with an increasing proportion of type 1 fibers in the deeper Type 1fibers have lower glycogen content cornpared to type II fi ber^,"^^ which would explain the differences in glycogen concentrations between the different samplings depths. The PG and MG concentration was higher in the more superficial parts of the gluteus medius muscle, however these differences were only
significant for PG (P<0.05). The rnean ratio of MG was approximately 40% of the total
glycogen concentration in the gizltez~~rnediz(s muscle regardless of sampling depth, In rat
muscle the ratio between PG and MG is constant between different muscle fiber types." It
is, however, not possible to draw any conclusions fiom our data regarding the relationship
between PG and MG for different fiber types in the horse as this information requires
rneasurement of the glycogen forms in single fibers.
In conclusion, the AC and MG+PG determinations give an accurate representation of
total glycogen in muscle biopsies fiom the horse and the two methods are comparable in
their precision. The present study demonstrates that equine skeletal muscle contains MG
and PG and that the two fiactions can be measured accurately in biopsy-sized sarnples.
The proportion of MG was not constant at different glycogen concentrations but rather increased with increasing total glycogen content.
3.5 ACKNOWLEDGEMENTS
This study was generously supported by EP Taylor Equine Trust Fund. We wouId like to thank the horse owners for allo.wing us to sarnple their horses.
"Minitab Inc. State College, PA, USA, 01997. O 1O0 200 300 400 500 600 700 AC (rnmol glucosyl unitslkg dw)
Figure 1. A cornparison of total muscle glycogen in horse muscle by using acid hydrolysis
(AC) and macroglycogen (MG) + proglycogen (PG) detemination (expenment 1). Each point represents an individual muscle biopsy. Solid line is linear regression analysis: y=x-9.2, r=0.99, n=45; dashed line, line of identity with dope 1; dotted line 95% prediction interval. 100 200 300 400 500 600 700 Mean value (MGfPG and AC) for each biopsy (mmol gIucosyl unitslkg dw)
Figure 2. Scatter diagram of the paired differences between methods for analysis of total glycogen (rnacroglycogen + proglycogen, MWGand acid hydrolysis, AC) against rnean of both data. Each point represents an individual muscle biopsy hmthe horses in experiment 1 (n=45). Solid Iine is mean difference between MG+PG and AC (-7.1 mol glucosyl unitskg dw); dotted line is limit of agreement, mean difference 2 SD ( -34.3 to
20.1 glucosy1 unitskg dw). O 50 100 150 200 250 300 MG (mm01 glucosyl units/kg dw)
Figure 3. A cornparison of macroglycogen (MG) concentrations and MG + proglycogen
(PG) concentrations obtained fkom individual muscle biopsies (n=31) fiom the middle gluteal muscle in expenment 1; y=(6.4904x)/(Z+O.O079x); r-0.90, P<0.000 1 Table 1: Reproducibility for total glycogen in duplicate analyses Erom the sarne muscle biopsy for the AC and MG+PG technique.
Measurement N Mean SD SE CV CR
MG Gly,,, c 400 rnrnoVkg dw 10 Gly,, > 400 mrnol/kg dw 35 Al1 sarnples 45 PG Gly,,, c 400 mmol/kg dw 10 Gly,,, > 400 mmoVkg dw 35 Al1 samples 45 MG+PG Gly,,, c 400 mrnolkg dw 10 Gly,,, > 400 mmoVkg dw 35 Al1 sarnples 45 AC Gly,, c 400 rnrnoVkg dw 10 Gly,,,, > 400 mrnoVkg dw 35 Al1 sarnples 45 454.2 14.6 0.7 3 -2 4 1.4 N, SD, SE, CV and CR represents the number of duplicates, standard deviation, standard error of the mean, coefficient of variation and coefficient of repeatability. Al1 values are in mrnoL%g dry weight (dw), except for coefficient of variation (CV), which is reported as a percent; MG, rnacroglycogen; PG, proglycogen, AC, acid hydrolysis; G~Y,~,total g1 ycogen. Table 2: Reproducability in duplicate biopsies obtained at the sarne site in the glzrtezts medilis muscle (9 horses). Mean Measurement Biopsy 1 Biopsy 2 SD SE CV CR
SD, SE, CV and CR represents the standard deviation, standard error, coefficient of variation and coefficient of repeatability. Al1 values are in mmoVkg dry weight (dw), except for coefficient of variation (CV), which is reported as a percent; MG, macroglycogen; PG, proglycogen.
Table 3 : The effect of sampling depth in the gluteus menitis muscle. Sampling depth Measurement 40 mm 80 mm
Al1 values are in mmoUkg dry weight (dw) * SE; MG, macroglycogen; PG, proglycogen. *Significantly different (Pc0.05) fiom sampling depth 40 mm. 3.6 REFERENCES
1. Bergstrom J. Muscle electrolytes in man. Determination by neutron activation
analysis on needle biopsy specimens. A study on normal subjects, kidney patients and
patients with chronic diarrhoea. Scand Jclin La6 lnvest 1962; 14:Suppl. 68
2. Kits Van Heijningen AJM, Kemp A. Free and fixed glycogen in rat muscle.
Bfocherri J l955;59:487-49 1.
3. Stetten Jr. DyStetten MR, Glycogen Metabolism. Physiol Rev 1960;40:505-537.
4. Jansson E. Acid soluble and insoluble glycogen in human skeletal muscle. Acta
Physiol Scand 1981;113:337-340.
5. Friden J, Seger J, Ekblom B. Topographical Iocalization of muscle glycogen: an ultrahistochemical study in the hurnan vastus lateralis. Acta Physiol Scand 1989; 135:3 8 1-
391.
6. Lomako J, Lomako WM, WheIan WJ, et al. Glycogen synthesis in the astrocyte: from glycogenin to proglycogen to glycogem FASEB J 1993;7: 1386- 1393.
7. Alonso MD, Lomako J, Lomako WM, et al, A new look at the biogenesis of glycogen. FASEB J 1995;9: 1 126-1 137.
8. Adarno KB, Graham TE. Comparison of traditional measurements with macroglycogen and proglycogen analysis of muscle glycogen. . J Appl Physiol l998;84:908-9 13. 9. Lomako J, Lomako WM, Whelan WJ. Proglycogen: a low-molecular-weight
form of muscle glycogen. FEBS Lett 199l;279:223-228.
10. Graham TE, Adarno KB, Shearer J, et al. Pro- and rnacroglycogenolysis:
relationship with exercise intensity and duration. JAppl Physiol2001;90:873-879.
Il. Hansen BF, Derave W, Jensen P, et al. No limiting role for glycogenin in
determining maximal attainable glycogen levels in rat skeletal muscle. Am J Physiol
Endocrinol Metab 2000;278:E398-404.
12. Shearer J, Marchand 1, Tamopolsky MA, et al. Pro- and macroglycogenoIysis
during repeated exercise: roles of glycogen content and phosphorylase activation. J Appl
Physiol2001;90:880-888.
13. Derave W, Gao S, Richter EA. Pro- and macroglycogenolysis in contracting rat
skeletal muscle. Acta Physiol Scand 2000; l69:29 1-296.
14. Adamo KB, Tamopolsky MA, Graham TE. Dietary carbohydrate and
postexercise synthesis of proglycogen and macroglycogen in human skeletal muscle. Am
J Physiol l998;27S:EZ29-334.
15. LindhoIm A, Piehl K. Fibre composition, enzyme activity and concentrations of metabolites and electrolytes in muscles of standardbred horses. Acta Vet Scarzd
1974;15:287-309.
16. Passonneau JV, Lauderdale VR. A Cornparison of three methods of glycogen measurement in tissues. Analytical Biochemist~1974;60:405-4l2. 17. Bland JM, Altman DG. Statistical methods for assessing agreement between two methods of clinical measurement, Lancet 1986; 1:3 07-3 1O.
18. Hanis RC, Hultman E, Nordesjo LO. Glycogen, glycolytic intermediates and high-energy phosphates determined in biopsy sampIes of rnuscuIus quadriceps femoris of man at rest. Methods and variance of values. Scand J Clin Lab hrvest 1974;33: 109- 120.
19. Snow DH, Baxter P, Rose W. Muscle fibre composition and glycogen depletion in horses cornpeting in an endurance ride. Vet Rec 198 1; 108:374-3 78.
20. Essen-Gustavsson B, McMiken D, Karlstr6m K, et al. Muscular adaption of horses during intensive training and detraining. Equine ver J l989;2 1:27-33.
2 1. Brammer GL, Rougvie MA, French D. Distribution of a-arny lase-resistent regions in the glycogen molecule. Carbohydr Res 1972;24:343-354.
22. Lomako J, Lornako WM, Whelan WJ. Glycogen metabolism in quai1 ernbryo muscle. The role of the glycogenin primer and the intemediate proglycogen. Eur J
Biocheizz 1995;234:343-349.
23. Roach PJ, Skurat AV. Self-glucosylating initiator proteins and their role in glycogen biosynthesis. Prog Nucleic Acid Res Mol Biol 1997;57:289-316.
24. Nevill ME, Boobis LH, Brooks S, et al. Effect of training on muscle metabolism during treadmill sprinting. J ApplPhysiol 1989;676:2376-2382. 25. Serrano AL, Petrie JL, Rivero JL, et al. Myosin isoforms and muscle fiber characteristics in equine gluteus medius muscle. Anat Rec l996;244:444-45 1.
26- Karlstrom K, Essén-Gustavsson B, Lindholm A. Fibre type distribution, capillarization and enzyrnatic profile of locomotor and nonIocomotor muscles of horses and steers. Acta Anatornica 1994;151:97-106.
27. Essen B, Henriksson J. Glycogen content of individual muscle fibres in man.
Acta Physiol Scarzd 1 974;90:645-647. CHAPTER 4
4.0 GENERAL DISCUSSION
In the present study, there was a strong positive correlation between total glycogen
measured by acid hydrolysis (AC) and by the proglycogen-macroglycogen (PG+MG)
determination technique over a wide range of glycogen concentrations using linear
regression analysis (y = x - 9.2; r = 0.99). Linear regression analysis- is a cornmon
approach to compare two measurement techniques but the correlation coefficient and the
regression analysis should be interpreted cautiously. The exclusive use of the correlation coefficient as a statistical tool in these situations cm be misleading and inappropriate for several reasons. The correlation coefficient is a rneasure of association and it would be surprising if two methods designed to rneasure the same thing were not related. The correlation coefficient depends on the range of measurements used and could therefore be increased by choosing widely spaced observations (Campbell and Machin 1993). It is more appropriate to analyze how well two measmernent techniques agree (BIand and
Altman 1986). The linear regression line in our study was almost identical to the line of identity; which indicated that the two analysis methods were likely to agree very well. We confirmed the agreement between the two methods by using an alternative statistical method proposed by Bland and Altman (1986). The limit of agreement was -34.2 to 20.2 mm01 glucosyl units per kg dry weight and these lirnits were srnaller than the lirnits of agreement for duplicate analysis for total glycogen obtained by analysis with the AC and
PG+MG deterrnination technique. Repeatability is also of importance in studies where two methods are compared
because the variation within each method limits the amount of agreement. If one method
has poor repeatability the agreement between the two methods will be poor (Bland and
Altman 1990)- Our results showed that the AC and PG+MG detennination techniques had
alrnost identical values for thé coefficient of repeatability (Chapter 3, Table 1). The within
and between biopsy variability for total glycogen gave CV values of approxirnately 4%
for total glycogen analyzed by the AC and PGtMG determination technique. The results
of repeatability (CV and SD; Chapter 3, Table 1) for total glycogen determined by AC and
the PWMG technique were in good agreement with previous studies (Hultman 1967,
Essen and He~ksson1974, Harris et al. 1974, Lindholm and Piehl 1974, Adarno and
Graham 1998). Together the linear regression equation, the lirnits of agreement and the
values of repeatability indicate a strong agreement between the AC and PG+MG
detemination techiques for total glycogen.
The present study evaluated repeatability between muscle biopsies exclusively fiom
restins horses. The variation between muscle biopsies would probabiy have been higher if
the study were conducted on post-exercise sarnples. During exercise the fiber recruitment varies within the muscle, which results in differences in the glycogen depletion pattern
(Essen 1978, Snow et al. 1981, Valberg 1986, Gottlieb 1989). This increase in the non- uniform distribution of glycogen in the skeletal muscle after exercise can potentially lead to higher variation in total glycogen for post-exercise sarnples. Considering this non- uniform activation of muscle fibers within the muscle the PGf-MG determination technique permits determination of total glycogen, proglycogen (PG), macroglycogen (MG) and muscle metabolites fiorn the same piece of tissue, which eliminates the
variation between muscle pieces.
Another purpose of this study was to determine whether the concentrations of PG and
MG are stable during the extraction procedure with regard to extraction time and PCA
concentration. If the relationship behveen the two forms of glycogen continues to change with extraction time the reliability of the separation technique must be questioned and the biological significance of the data would be limited. Our results showed that the concentrations of PG and MG were stable with extraction time. The stability seems to be more pronounced for extraction in 0.5 M and 1.5 M PCA cornpared to extraction in 3.0 M
PCA (Chapter 2, tables 1-3). Extraction in 3.0 M PCA yielded lower PG values and higher
MG concentrations (%O.OS) for al1 studied extraction tirnes (10-40 minutes). The possibility that stronger acid concentration yielded higher MG concentrations due to pre- hydrolysis of glycogen into glucose residues by the PCA was excluded by analysis of glucose in the PCA extracts prior to acid hydrolysis in HC1. The mean glucose concentration in the PCA extracts for each of the treatment combinations of different PCA concentrations and extraction times were almost identical (range 0.45 - 0.54 mm01 glucose per kg dry weight). Instead it is possible that the stronger acid (PCA 3.0 M) caused a limited arnount of hydrolysis of small chains of polysaccharides from the PG fiaction, which joined the PCA-soluble fraction (MG fiaction). This theory is supported by the fact that the change in the PG and MG pool tend to be inversely related.
A problem with the PG+MG detemination technique is that endogenous tissue glucose and glucose-6-phosphate (G6P) are recovered in the MG fraction and these metabolites will therefore be incorrectly measured as MG with the hexokinase method. In resting muscle the concentrations of these metabolites are very small relative to the MG
fiaction and they couid therefore be ignored. However, under conditions of intense
exercise the intracellular concentration of glucose and G6P increases, which could cause a
potential error in the measured MG fiaction (Stetten et al. 1958, Derave et al. 2000).
When the glucose residues are analyzed after HC1 hydrdysis with the hexokinase method
according to Bergrneyer (Bergmeyer 1974) the steps in the analysis method are perforrned
in such a way that the G6P concentration will be eliminated from the MG fiaction because
the enzyme hexokinase is not added until after the initial reading of NADPH. It is,
however, crucial that G6P is stable during the acid hydrolysis of glycogen. We recently
showed that the concentration of G6P is stable after boiling G6P in 1 M HCl for up to 2
hours (unpublished observations, Brojer and Essen-Gustavsson). Separate analysis of
intracellular glucose is thus required for post-exercise samples in order to obtain accurate concentrations of MG. The post exercise sarnples in our study, however, were al1 taken several hours after exercise and are therefore assumed to contain values of intracellular glucose similar to resting values.
Previous studies conducted on human and rat muscle have dernonstrated that the relative proportions of PG and MG on a weight basis were dependent on the total glycogen concentration (Adarno and Graham 1998, Adamo et al. 1998, Hansen et al.
2000). As total glycogen concentration increased in the muscle the proportion of MG increased whereas the proportion of PG decreased. Despite a decrease in the relative proportion of glycogen, the absolute PG concentration increased as total glycogen increased. A similar relationship has been demonstrated in this study for equine skeletal muscle (Chapter 3, figure 3). It appears £rom previous studies in humans that the concentration of PG has a
maximal concentration of approximately 250-300 mm01 glucosyl units per kg dw in
skeletal muscle, With increasing total glycogen concentrations the PG concentration
increases up to 250-300 molglucosyl units per kg dw, which occurs at a total glycogen
concentration of approximately 300-350 mm01 glucosyl units per kg dw. Increase in total
glycogen over this concentration appears to be a result of increase in the MG pool. This
threshold of 350 mm01 glucosyl units per kg dw corresponds to the normal resting
concentration of muscle glycogen in human skeletal muscle. This pattern was not
identified for equine skeletal muscle in this study as the PG concentration continued to
increase up to 350 mm01 glucosyl units per kg dw. It is possible that PG reaches a plateau
at higher glycogen concentrations but to study this hypothesis muscle biopsies with higher
total glycogen concentrations than those available in this study are required.
There is controversial information in the literature regarding whether PG and MG
exist as distinct molecular sizes or as a range of sizes (Lomako et al. 1993, Alonso et al.
1995, Lomako et al. 1995, Roach and Skurat 1997). Previous studies in rat and human
muscle have indicated that the average sizes of the PG molecule increases with increasing total glycogen-levels (Adamo et al. 1998, Hansen et al. 2000). This issue could not be addressed in our study even though the absolute PG concentration increased with increasing total glycogen levels because al1 the muscle biopsies included in our study were taken from different individuals or from different locations (gluteal muscle vs triceps muscle). The results fi-orn this study also illustrate the importance of a standardized sarnpling site and depth in the muscle. In the gluteal muscle the total glycogen, PG and
MG concentrations increased with increased sarnpling depth (statistically~significantfor G, and PG, but not for MG; PcO.05). This is most likely a result of an increasing proportion of type I fibers with increasing depth in the gluteus medius muscle (Karlstrom et al. 1994, Serrano et al. 1996). The relationship between the PG and MG concentrations was almost identical between the two different sampling depths. It is however not possible to draw any conclusions firom Our data regarding the relationship between PG and MG for different fiber types in the horse. This information requires rneasurement of PG and MG in single fibers. Interestingly, studies in rat skeletal muscle have identified a similar relationship between PG and MG regardless of the fiber type (Hansen et al. 2000).
4.1 GENERAL CONCLUSIONS
In summary, this study showed the existence of PG and MG in equine skeletaI muscle and validated a technique for analysis of these two fiactions of glycogen in fireeze dried muscle sarnples fiom the horse. Both the AC and the PG+MG determination technique provided good, repeatable measures of total glycogen in fieeze dned muscle biopsies fiom the horse. The agreement between the two methods for analysis of total glycogen was also very good. The two pools of glycogen were very stable with extraction tirne and concentration of PCA in the range between 0.5 and 1.5 M during the separation procedure.
Extraction in 3.0 M PCA yielded signïficantly different values for PG and MG and this acid concentration was therefore not considered to be usehl for separation of the two glycogen pools in equine skeletal muscle. Further, the biopsy depth was shown to have an effect on the concentration of the two pools of glycogen and it was therefore considered important to standardize the site and depth for the biopsies. The relative proportions of PG and MG on a weight basis were dependent on the total glycogen concentration. As total glycogen concentration increased in the muscle the proportion of MG increased whereas the proportion of PG decreased. Despite a decrease in the relative proportion of glycogen, the absolute PG concentration increased as total glycogen increased. The PG+MG deterrnination technique permits determination of total glycogen, PG, MG and muscle metabolites from the same piece of tissue, which eliminates the variation between muscle pieces. This shidy is the first to document PG and MG in the equine skeletal muscle.
4.1 FUTURE STUDIES
The development of a good reliable method for determination of PG and MG in equine skeletal muscle should enable investigation of the physiological response of glycogenolysis, synthesis and regulation of the two fractions of glycogen. To date very limited information is available regarding the physiological fùnction and the metabolic differences between PG and MG.
In resting human muscle 6575% of the glycogen on a weight basis is in the PG fonn
(Adamo and Graham 1998, Adamo et al. 1998, Asp et al. 1999). Adamo and coworkers
(1998) studied the re-synthesis of PG and MG in human skeletal muscle after glycogen depleting exercise. They found that PG was the most dynamic form of glycogen during recovery fiom exercise and that the majority of the glycogen synthesized over the first 4 hours was in the PG pool. In another study Adamo and Graham (1998) dernonstrated that both PG and MG were used as substrates for glycogenolysis in human skeletal muscle but the net rate of the glycogenolysis was higher for the PG pool. Glycogen is found in higher quantities in equine skeletal muscle than in other species (Snow and Valberg 1994). In the gluteal muscle of untrained horses approxirnately 60% of the total glycogen is in the PG
77 form. In well trained horses with total glycogen concentrations of over 600 molglucosyl
units per kg dw the percentage of PG appears to be even Lower (unpublished data Brojer
2001). The repletion of glycogen to baseline values appears to occur relatively slowly in
the horse compared to hurnans and rats (Snow et al. 1987). Due to the differences between
horses and man it is possible that a different pattern exists in the horse regarding re-
synthesis and glycogenolysis for the two different pools of glycogen and it is therefore
important to address these questions in füture studies.
In equine skeletal muscle the glycogen content and the glycolytic capacity between muscle fibers varies; type 1 fibers have lower glycogen content and gIycolytic capacity than type JI fibers (Snow et al. 1981, Valberg 1986). The nurnber and fiber types that are recruited during exercise are dependent on type, speed and duration of exercise. In general the fiber recruitment follow the order fkom type 1 through type IIA to type ID3 fibers
(Essén-Gustavsson et al. 1984, Valberg 1986) (White and Snow 1987, Gottlieb 1989).
Knowing that PG and MG differ in their metabolic regulation and that the fiber types differ with regard to glycogen content, glycolytic capacity and fiber recruitment one would expect differences in the PG and MG concentrations between fiber types. In the rat skeletal muscle PG and MG content varies between fiber types but the ratio between PG and MG is similar (Hansen et al. 2000). It is, however, possible that species differences exist and it is therefore important to determine whether differences in PG and MG content exist within fiber types in the horse. Future studies on single fibers could be used to increase the understanding of the metabolism of these two pools of glycogen. It is crucial to address the implications of the two different foms of glycogen for the different equine athletes; standardbred, thoroughbred and endurance ride horses. A number of horses with chronic exertional rhabdomyolysis have recently been found to have a glycogen storage disorder characterized by the accumulation of an abnormal polysaccharide in the type DA and type IIi3 fibers (polysaccharide storage myopathy;
PPSM) (Valberg et al. 1997, Valentine et al. 1997). The abnormal polysaccharide is very slow to digest with amylase. The total glycogen content in the skeIetal muscle is 1.5 - 4 times higher in horses with PPSM compared to normal horses (Valberg et al. 1997).
Horses with PPSM can generate Iactic acid during exercise and al1 of the activities of glycolytic enzymes are normal indicating that the glycogen accumulation is not caused by an inability to metabolize glycogen. Rather, horses with PSSM appear to have a novel defect characterized by enhanced glucose storage and glycogen synthesis (Valberg et ai.
1999, Valberg et al. 1999). It is possible that studies of the metabolism of PG and MG in horses with PSSM would develop curent pathophysiologic understanding of the diseases. 4.3 REFERENCE LIST
Adamo, K.B. and Graham, T.E. (1998) Cornparison of traditional measurements with
macroglycogen and proglycogen analysis of muscle glycogen. J Appl Physiol- 84,
908-913.
Adamo, K.B., Tarnopolsky, M.A. and Graham, T.E. (1998) Dietary carbohydrate and
postexercise synthesis of proglycogen and macroglycogen in human skeletal
muscle. Am J Ph-vsiol. 275, E229-334.
Alonso, M.D., Lomako, J., Lomako, W.M. and Whelan, W.J. (1995) A new look at the
biogenesis of glycogen. FASEB J. 9, 1126-1 137.
Asp, S., Daugaard, J.R., Rohde, T., Adamo, K. and Graham, T.E. (1999) Muscle
glycogen accumulation after a marathon: roles of fiber type and pro- and
macroglycogen. JAppl Physiol. 86,474-478.
Bergmeyer, H. (1974) Methods of Enzymatic Analysis, Academic, New York. vol 3, pp
1128-1 131.
Bland, J.M. and Altman, D.G. (1986) Statistical methods for assessing agreement
between two methods of dinical measurement. Lancet. 1, 307-3 10.
Bland, J.M. and Altman, D.G. (1990) A note on the use of the intraclass correlation
coefficient in the evaluation of agreement between two methods of rneasurement.
Contput Biol Med. 20,337-340. CampbeI1, M.J. and Machin, D. (1993) Common pitfalls in medical statistics. In: Medical
Statistics a Cornmonsense Approach, 2nd edn. John Wiley & Sons, Chichester. pp
126- 136,
Derave, W., Gao, S. and Richter, E.A. (2000) Pro- and macroglycogenoIysis in
contracting rat skeletal muscle. Acta Physiol Scand. 169,291-296.
Essen, B. (1978) Glycogen depletion of different fibre types in human skeletal muscle
during intermittent and continuous exercise- Acta Physiol Scand. 103,446-455.
Essen, B. and He~ksson,J. (1974) Glycogen content of individual muscle fibres in man.
Acta Physiol Scand. 90, 645-647.
Essen-Gustavsson, B., Karlstrom, K. and Lindholrn, A. (1984) Fiber type, enzyme
activities and substrate utilization in skeletal muscle of horses competing in
endurance rides. Equine Vet J. 14, 197-202.
Gottlieb, M. (1989) Muscle glycogen depletion patterns during draught work in
Standardbred horses. Equine Vet J. 21, 110-1 15.
Hansen, B.F., Derave, W., Jensen, P. and Richter, E.A. (2000) No limiting role for
glycogenin in determinhg maximal attainable glycogen Ievels in rat skeletal
muscle. Am J Physiol Endocrinol Metab. 278, E398-404. Harris, R.C., Hultman, E. and Nordesjo, L.O. (1974) Glycogen, glycolytic intermediates
and high-energy phosphates determined in biopsy samples of muscuIus quadriceps
femoris of man at rest. Methods and variance of values. Scand J Clin-LabInvest.
33, 109-120.
Hultman, E. (1967) Muscle glycogen in man determined in needle biopsy specimens:
method and normal values. Scand J Clin Lab Invest. 19, 209-217.
Karlstrom, K., Essén-Gustavsson, B. and Lindholm, A. (1994) Fibre type distribution,
capillarization and enzymatic profile of locomotor and nonlocornotor muscles of
horses and steers. Acta Arzat (Basel), 151, 97-1 06.
Lindholm, A. and PiehI, K. (1974) Fibre composition, enzyme activity and concentrations
of metabolites and electrolytes in muscles of standardbred horses. Acta Vet Scand
15,287-309.
Lomako, J., Lomako, W.M. and Whelan, W.J. (1995) Glycogen metabolism in quail
embryo muscle. The role of the glycogenin primer and the intermediate
proglycogen. Eur J Biochem. 234, 343-349.
Lomako, J., Lomako, W.M., Whelan, W.J., Dombro, R.S.,Neary, J.T. and Norenberg,
M.D. (1993) Glycogen synthesis in the astrocyte: fiom glycogenin to proglycogen
to glycogen. FASEB J. 7, 1386-1393.
Roach, P.J. and Skurat, A.V. (1997) Self-glucosylating initiator proteins and their role in
glycogen biosynthesis. Prog Nztcleic Acid Res Mol Biol. 57, 289-3 16. Serrano, A.L., Petrie, J.L., Rivero, J-L, and Hermanson, J.W. (1996) Myosin isoforms
and muscle fiber characteristics in equine gluteus medius muscle. Anat Rec. 244,
444-45 1.
Snow, D.H., Baxter, P. and Rose, R.J. (1981) Muscle fibre composition and glycogen
depletion in horses competing in an endurance ride. Vet Rec. 108,374-378.
Snow, D.H., Harris, R.C., Harman, J-C. and Marlin, D.I. (1987) Glycogen repletion
following different diets. In: Equine Exercise Physioloay 2, Eds: SR Gillespie and
NE Robinsson. ICEEP Publication, Davis, California. pp 70 1-710.
Snow, D.H. and Valberg, S.J. (1994) Muscle anatomy, physiology, and adaptions to
exercise and training. In: The Athletic Horse: Princ@les and Practice of Equine
Medicine, Eds: DR Hodgson and RJ Rose. W. B. Saunders Company,
P hiladelphia. pp 145- 179.
Stetten, M., Katzen, H. and Stetton, D. (1958) A cornparison of the glycogen isolated by
acid and alkaline procedure. JBiol Chem. 232,475-488.
Valberg, S. (1986) Glycogen depletion patterns in the muscle of standardbred trotters after
exercise of varying intensities and durations. Equine Vet J. 18,479-484.
Valberg, S.J., MacLeay, J.M., Billstrom, J.A., Hower-Moritz, M.A. and Mickelson, J.R.
(1999) Skeietal muscle metabolic response to exercise in horses with "tying-up"
due to polysaccharide storage myopathy. Equine Vet J, 31,43-47. Valberg, S.J., MacLeay, J.M. and Mlckelson, J.R. (1997) Polysaccharide storage
myopathy associated with exertional rhabdomyolysis in horses. Comp Cont Educ
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Valberg, S.J., Mickelson, J.R., Gallant, E.M., MacLeay, J.M., Lentz, L. and De La Corte,
F. (1999) Exertional rhabdomyolysis in quarter horses and thoroughbreds : one
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Valentine, B.A., CrediUe, KM., Lavoie, J.P., Fatone, S., Guard, C., Curnrnings, J.F. and
Cooper, B.J. (1997) Severe polysaccharide storage rnyopathy in Belgian and
percheron draught horses- Eqzrine Vet J. 29,220-225.
White, M.G. and Snow, D.H. (1987) Quantitative histochemical study of glycogen
depletion in the rnaximaIIy exercised thoroughbred. Equine Vet J. 19, 67-69. CHAPTER 5
5.0 MASTER REFERENCE LIST
Adarno, K.B. and Graham, T.E. (1998) Cornparison of traditional measurements with
rnacroglycogen and proglycogen analysis of muscle glycogen. J Appl Physiol. 84,
908-9 13.
Adamo, K.B., Tarnopolsky, M.A. and Graham, T.E. (1998) Dietary carbohydrate and
postexercise synthesis of proglycogen and macroglycogen in human skeletal
muscle. Am J Physiol. 275, E229-334.
Alonso, M., Lagzdins, E., Lornako, J., Lornako, W. and Whelan, W. (1995a) New and
specific nucleoside diphosphate glucose substrates for glycogenin. FEBS Lett. 359,
110-1 12.
Alonso, M.D., Lomako, J., Lomako, W.M. and Whelan, W.J. (1995b) A new look at the
biogenesis of glycogen. FASEB J. 9, 1126-1 137.
Asp, S., Daugaard, J.R., Rohde, T., Adarno, K. and Graham, T.E. (1999) Muscle
glycogen accumulation after a marathon: roles of fiber type and pro- and
macroglycogen. J Appl Physiol. 86,474-478.
Bergmeyer, H. (1974) Methods of Enzymatic Analysis, Academic, New York. vol 3, pp
1128-1131. Bergstrom, J. (1962) Muscle electrolytes in man. Determination by neutron activation
analysis on needle biopsy specirnens. A study on normal subjects, kidney patients
and patients with chronic diarrhoea. Scarzd J ChLab Invest. 14, Suppl. 68.
Bergstrom, J., Hermansen, L., Hultman, E. and Saltin, B. (1967) Diet, muscle glycogen
and physical performance. Acta Physiol Scnnd 71, 140- 150.
Bland, J.M. and Altman, D.G. (1986) Statistical methods for assessing agreement
between two methods of clinical measurement. Lancet. 1, 307-3 10.
Bland, J.M. and Altman, D.G. (1990) A note on the use of the intraclass correlation
coefficient in the evaluation of agreement between two methods of rneasurement.
Cornput Bi01 Med. 20, 337-340.
Bramrner, G.L., Rougvie, M.A. and French, D. (1972) Distribution of' a-arnylase-
resistent regions in the glycogen molecule. Curbohydr Res. 24, 343-354.
Brojer, 1. T. (2001) Proglycogen and macroglycogen in equine skeletal -muscle. [MSc
Thesis], University of Guelph, Guelph, Ontario.
Calder, P.C. (199 1) Glycogen structure and biogenesis. Int J Biochem. 23, 1335- 1352.
Campbell, D.G. and Cohen, P. (1989) The amino acid sequence of rabbit skeletal muscle
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Table A: 1. Individual data for proglycogen (PG), rnacroglycogen (MG), PG+MG and acid hydrolysis (AC) fiom 45 individual muscle biopsies. Subject MuscIe Duplicate PG MG PG+MG AC a gluteal 1 31 /.4 128.2 505.6 51 5.6 2 360.6 136.1 496.7 51 8.9 a triceps 1 3X3Tg9- 108.4- 492T3-53TT9- 2 367.7 106.5 474.2 514.2 b gluteal 1 3-593-C4375-505:5-5i3:2- 2 367.1 152.8 51 9.8 51 5.2 b triceps 1 35B78p1-23T5-487:3-509T8- 2 364.3 135.7 500.1 508.9 c gIutestl 1 3-68T662110T00 57BT66589T33 2 373.9 196.3 570 -2 569.9 c triceps 1 3-119T221Q33 462T3-497Y7- 2 336.3 165.3 501 -5 500.1 d g IÜteai 1 326T2- 2T2-537:7539:4- 2 324.1 21 3.2 537.3 548-4 3-330T9p-- -- - d triceps 1 110.4 44T3- 47 7.1- 2 324.7 1'i 5.8 440.5 444.6 e gIüEdp- 1 3-5iT0p273T7p562:8p 526T0° 2 346.3 192.1 538.3 561 -6
e triceps 1 376.1 104.7 480r8 489T6 - 2 372.6 110.8 483.3 51 0.7 f triceps 1 3T6T2- 83T7-402T993358T55 2 340.7 87.3 428.0 43 1-4 f gluteal 1 301.4762TO 463T4- 5-- 18T3- 2 323.4 208.6 532 500.8 g gluteal 1 328T9924BT55 5753-58-971- 2 333.7 248.1 581 -9 598.2 9 triceps 1 33-975-1-5 9T679-93p5X5- 2 348 -3 153.9 502.2 521 -5 h triceps 1 296T0° 4g.1 333737ïT 2 281 .O 41 -3 322.2 359.0 I gluteal 1 24210-23276p 50X664722- 2 271 .O 201 -1 472.1 499.1 j triceps 1 38577725T8- 5'-iC5---7 OTO" 2 355.9 120.3 476 -2 483.1 1 gluteat 1 3-3iTOT53T444B4T4.491TO- 2 339.7 141.8 48 1.5 496.4 k gluteal 1 503T22272T66 575T8- 549Y9- 2 284.7 274.6 559.3 571 -6 I gluteal 1 193T8 10-5393000T7-299T7-'- 2 197.9 110.1 308.0 306.1 - - m gluteai 1 1922 -- 6 7r6- 259T8827274- 2 198.4 72.0 270.4 290 -8 n gluteal 1 23633763T84023 41 23- 2 260.2 183.6 443.8 41 6.5 Table A: 1 continued. O gluteal 1 106 26.5 132-5 152.2 2 120.2 24.9 145.1 158.6 P gIUteaI----I 2a1.1 111.3p 372-4- 3Z5- 2 243.3 132.5 375.7 364.6 9 gluteal 1 2277 76 297:8-292:8- 2 21 7.1 78.1 295.2 325.3 r gluteal 1 237- f6-33- 403, 1 478T7- 2 242.4 164.3 406.6 393.6 s g 1x1 1 277:s-273T6-9 1-1 5ZOTZ2 2 306.7 21 1.7 51 8.4 51 6.9 s triceps 1 3-31 -1 147T277BT3-47OT4- 2 336.5 135.8 472.3 452.1 t g lüiëZl 252.3- 92.4 3X77324T7p 2 238 -3 98.7 337 360.5 t triceps 1- 267 44.6- 3x6-329T 2 267.8 48.9 31 6.6 31 7.6 u glu teali276y6- f27:5-- 404.1 43-83- 2 250 145.5 395.5 428.4 u triceps 1 3ai.4-53. 1 354Zi537577- 2 304.5 53.4 358 370.5 v gluteal 1 314.1 10775 4 1-5T6-4-C 2 3 14.1 130.3 444.5 431 -3 v triceps 1 343.1 72T6-617- 3-97r8- 2 333.5 66 399.5 41 7.7 x glÜm 1 312198 182T3- 572.1 456T3- 2 312 180.6 492.6 493.2 x triceps 1 3aa7T6 1 1578p4-6-3T3p470ï4- 2 337.3 120.7 458 468.1 Y gluteal 1 274.1 2233.1 49x2-53-3T9- 2 281 -5 21 7-2 498.6 524-1 - - z giütëai - -- i--- 3 28:9--- 197T6p526Xp 5283- 2 372.9 186.6 559.5 547.4 aa glüfëd 1 26479 15C9- H9:8-437:5- 2 269.5 149.9 41 9.4 436.5 bb gluteal 1 3a3T6 1 19T5-4233429T6- 2 314.7 1 19.3 434 428.9 CC glütë5l 1 236TZ2256. 1 542T3352?.1- 2 271 254.5 525.4 545.7 -da gluteal 1 226 1-55T8-3878- 318115- 2 228.1 158.5 386.6 382.1 ee gluteal 1 272T5236.4508.9541.3- 2 281 -3 261 -1 542.3 507.7 ff gluteal 1 25378p23T97S3T6p483T7- 2 273.6 206.7 480.3 473 gg gluteal 1 307y66219T6- 5'27r2-54714 2 314.1 223.2 537.3 548.1 Values are reported as mm01 glucosyl units per kg &y weight. Table A:2. Individual data for proglycogen (PG),macroglycogen (MG) and PGtMG fiom biopsies obtained at different sarnpling depths in the middle gluteal muscle. 40 mm Sampling Depth 80 mm Sampling Depth Subiect PG MG PG+MG PG MG PG+MG 1 285.6 190.0 475.6 255.8 203.2 458.9 2 28 1.8 204.3 486.1 259.4 202.2 461-6 3 273.9 144.8 418.7 258 -9 154.1 41 3.0 4 243 -6 1 14.0 357.7 235.8 101.O 336.8 5 278.9 29 1 -8 570.7 243 -7 272.1 515.8 6 271 -8 166.8 455.1 257.0 144.1 423.9 7 312.0 207.5 493.8 292.8 183.3 500.3 8 300.6 184.0 520 -7 271 -1 181-8 455.1 9 307.2 215.3 522.5 327.8 220.1 543.1 Values are reported as mm01 glucosyl units per kg dry weight, Table A:3. Individual data for proglycogen (PG), macroglycogen (MG) and PWMG fiom repeated biopsies obtained fkom the sarne site in the rniddle gluteal muscle- Sampling 1 Sampling 2 Su bject PG MG PG+MG PG MG PG+MG 1 2f 1.8 21 8.7 496-4 345.1 1 l8-l 523.8 2 350.9 192.1 543.0 305.5 189.4 494.9 3 267.2 1 52.4 41 9.6 274.6 129.3 403.9 4 309.2 119.4 428.5 309.1 103.6 41 2.7 5 278.6 255.3 533.9 268.2 299.8 569.7 6 227-1 157.1 384.2 231 -9 154.1 386.0 7 276 -9 248-7 525.6 273.5 267.2 540.6 8 263.7 21 0.8 474.5 281 -6 229.4 51 1 .O 9 31 0.8 221 -4 532.2 302.0 196.5 498.5 Values are reported as mm01 glucosyl units per kg dry weight. Table A:4. Individual data for progIycogen (PG), macroglycogen (MG) and PG+MG for different extraction tirnes. Extraction Subject Time PG MG PG+MG I 10 3l1.1 164.4 535.5
Values are reported as mm01 glucosyl units per kg dry weight. Table A5Individual data for proglycogen for different combinations of extraction times and perchloric acid (PCA) concentrations, Extraction PCA Subject Duplicate Time 0.5 M 1.5 M 3.0 M A 1 10 31t.3 365.9 345.8 Table A:5 continued. A 1 40 3 (9.8 J f 6.8 318.8 2 40 376.7 351.3 340.8 B 1 40 320.9 310.0 247.1 2 40 32 9.5 280.5 269 -7 C 1 40 268.2 272.8 256.1 2 40 271 -3 286.2 262.3 D 1 40 267.2 261 -3 21 5.9 2 40 261 -6 288.5 21 7.9 E 1 40 333.1 350.8 31 0.5 2 40 333.5 329.9 276.7 F 1 40 305.8 325.1 333.5 2 40 322.8 327.4 330.1 Values are reported as mm01 glucosyl units per kg dry weight. Extraction times are reported in minutes. Table A: 6. Individual data for macroglycogen for different combinations of extraction tirnes and perchloric acid (PCA) concentrations. Extraction PCA Subject Duplicate Time 0.5 M 1.5 M 3.0 M A 1 10 149.3 181.3 202.8 Table A:6 continued. A 1 40 119-1 1 14.2 230. 1 2 40 181-1 187.7 238.5 8 1 40 202-7 260.2 287.2 2 40 21OS 240.7 307-1 C 1 40 80.9 81 -5 120.4 2 40 81.5 94.0 121-9 D 1 40 158.7 154.3 191.7 2 40 145.7 153.5 209.1 E 1 40 209.7 182.4 268.9 2 40 182.4 204.4 235.6 F 1 40 39.0 33.4 37-1 2 40 36.8 31.3 34.6 Values are reported as mm01 glucosyl units per kg dry weight. Extraction times are reported in minutes. Table A: 7. Individual data for proglycogen + macroglyco gen for different combinations of extraction times and perchloric acid (PCA) concentrations. Extraction PCA Subject Duplicate Time 0.5 M 1.5 M 3.0 M A 1 10 526.5 / -2 548.6 Table A:7 continued. A 1 400 2 40 557.8 539 .O 579.3 B 1 40 523.6 570.1 534.3 2 40 530.0 521-1 576.7 C 1 40 349.1 354.4 376.6 2 40 352.8 380.2 384.2 D 1 40 425.8 41 5.7 407.6 2 40 407.3 442. 1 427.0 E 1 40 542.8 533.3 579.4 2 40 516.0 534-3 512.3 F 1 40 344.9 358.5 370.6 2 40 359.5 358 -7 364.8 Values are reported as mm01 glucosyl units per kg dry weight. Extraction times are reported in minutes. Table A:8. Individual data for glucose concentration in aliquots f?om the MG &actions before hydrolysis in HCI. Extraction PCA Subject Duplicate Time 0.5 M 1.5 M 3.0 M A 1 10 0.36 O -40 0-41 Table A:8 continued. A 1 40 0.56 0.51 0.5/-- 2 40 0.53 0.68 0 -70 B 1 40 O .44 0 -46 0 -46 2 40 0 -42 0 -46 0 -45 C 1 40 0.59 0.65 0-71 2 40 0.64 O -64 O -63 D 1 40 0.48 0.51 0.54 2 40 0 -52 0.52 0.53 E 1 40 0.49 0.29 0.1 8 2 40 0.47 0.60 0.14 F 1 40 0.59 O -49 0.65 2 40 0 -68 0.61 0.79 Values are reported as molglucose per kg dry weight, Extraction times are reported in minutes. 5.2 APPENDIX B
Pro- and Macroglycogen separation
1. Solutions
2 M Trizma Base: 60.55g Trizma Base (Boehringer Mannheim 604203) in 250 ml Milli-Q HzO-
1.5 M PCA: 3 1.5 ml PCA (70%) in 250 ml Milli-Q HzOI
2. Muscle
a) Freeze-dry and powder muscle. Great care must be taken to remove blood, connective tissue and extra-muscular fat. b) Weigh muscle directly into tared 5 ml pyrex tubes. The muscle weight should be between 1.5 - 3.0 mg. Store with dessicant until ready to proceed.
3. Extraction in PCA (separation of pro- and macroglycogen) a) Place the pyrex tubes with the muscle sarnples in a rack on a bed of ice. b) Add 200pl ice-cooled PCA to each sarnple. Mix samples with the PCA using a plastic rod. Extraction is to occur on ice for 20 minutes. c) Centrifuge at 3000 rprn for 15 minutes. d) Remove lOOpl of the PCA supernatant (macroglycogen fraction) and transfer the volume to a 5 ml pyrex tube. e) Aspirate the remaining PCA and keep the pellet (proglycogen fraction).
4. Hydrolysis of macroglycogen and proglycogen into glucose residues a) Add L ml of 1 M HCl to each of the pyrex tubes with proglycogen and rnacrogIycogen. b) Add caps to the tubes and record the weight. c) Boil the tubes in a wzter bath at ZOOOC for 2 hours.
d) Re-weigh the tubes and rnake up differences with distilled HzO if over 50 pi difference,
e) Neutralize aIl tubes with 1 ml of 2 M Trima Base.
f) Vortex and centrifuge at 3000 rpm for 5 minutes.
g) Transfer supernatant into eppendorf tubes and store at -80°C until analysis of glucosyl units.
Analysis of glucose units
1. Principle Glucose + Mg-ATP --- b Glucose-6-Phosphate + Mg-ADP
Measure the fluorescence of NADPH which is representative of glucose units.
2. Reagents
Triethano lamine Sigma T-1502 Magnesium C hloride Fisher 780247 ATP Boehringer 5 19 987 NADP Boehringer 128 059 or 128 040 G6PDH Boehringer 127 655 Hexokinase Boehringer 127 809 or 1426 362 Glucose Standard Sigma 635-100
3. Preparation of buffer
Amount of buffer needed is 1.O ml per tube. Mix together and stir well:
Buffer volume 50 ml 100 ml 150 ml 200 ml 300 ml Triethanolamine 2.8 g 5.6 g 8.4 g 11.2 g 16.8 g MgCl2 0.3 g 0.6 g 0.9 g 1.2 g 1.8 g Add Milli-Q H20 25 ml 50 ml 75 ml 100 ml 250 ml BM~to pH 7.5 using 2 M NaOH, then add required amount of ATP, NADP and G6PDH and bring up to final volume in graduated cyiinder (see below).
Buffer volume 50 ml 100 ml 150 ml 200 ml 300 ml ATP 80 mg 160 mg 240 mg 320 mg 480 mg NADP 10 mg 20 mg 30 mg 40 mg 60 mg G6PDH 2 Fl 4 PI 6 8 PI 12 pl
When buffer is complete stir well and cover.
Preparation of enzyme
G6PDH is added directly to buffer (see above),
Cdculate the arnount of hexokinase needed (10 putube).
Dilute hexokinase with Milli-Q H20, 1:9.
Mix the enzyme gently; do not vortex. Store in fndge.
Preparation of precinorm control
Dilute precinorm control with Milli-Q H20, 15(50 pl precinorm + 250 pl HzO). Concentration range for control: 5.15 - 6.29 mM.
Vortex and place in standard curve rack.
Preparation of standard curve
Concentration Milli-Q H20 Glucose standard 214 pM 500 pL 20 pL 412 FM 500 pL 40 pL 596 jA4 500 pL 60 PL 767 ph4 500 pL 80 a 927 pM 500 @ 100 pL 1854 p.M 400 pL 200 pL a) Use 12 / 75 tubes. b) Add the stock (glucose standard) solution according to the chart above. c) Add the required volume of MilIi-Q H20according to the chart. d) Vortex al1 standard solutions and place in standard curve rack. e) Pipette 20 pL of Milli-Q HzO in triplicates in the blanks (10 / 75 tubes) of the standard rack. f) Pipette 20 pL of standard solution in triplicates in the standard tubes (10 / 75 tubes) of the standard rack. g) Pipette 20 pL of control in triplicates in the control tubes (10 1 75 tubes) of the standard rack. h) Add 1.O rnL of buffer to each tube in the standard rack. i) Vortex al1 tubes in the standard rack.
7. Preparation of proglycogen and macroglycogen samples a) Prepare a mixed solution of 1 ml 1 M HC1 and 1 ml 2 M Trima-Base and vortex. b) Pipette 20 yS. of the mixed HCl and Trima-Base solution in tnplicates to the blanks on the sample racks (10 / 75 tubes). c) Pipette 20 $ of sarnple in triplicates in the sample tubes (10 / 75 tubes). d) Pipette 1.O mL of buffer to each 10 1 75 tubes on the sarnple rack (sarnples and blanks). e) Vortex al1 tubes in the sarnple racks.
8. Determination of gIucose units on fluorometer (LS-50) a) Read each tube on fluororneter (excitation 340, emission 455). b) Add 10 PL of enzyme (hexokinase) to each tube and vortex. c) Let stand for 1 hour, then take second reading on fluorometer.