REGULATION OF PROTEIN TURNOVER DURING HYPER-OSMOTIC
STRESS IN SKELETAL MUSCLE
Cody Vandommele, BKin.
Submitted in partial fulfillment of the requirements for the degree
Masters of Science in Applied Health Science
(Kinesiology)
Supervisor: Dr. Brian D. Roy
Faculty of Applied Health Sciences
Brock University
St. Catharines, Ontario
Cody Vandommele © January 2012
ABSTRACT
REGULATION OF PROTEIN TURNOVER DURING HYPER-OSMOTIC STRESS IN
SKELETAL MUSCLE
Cody Vandommele
Advisor Brock University
Dr. B.D. Roy
The purpose of this study was to examine the effect of hyper-osmotic stress on
protein turnover in skeletal muscle tissue using an established in-vitro model. Rat
EDL muscles were incubated in either hyper-osmotic (400 ± 10 Osm) or isoosmotic (290 ± 10 Osm) custom-modified media (Gibco). L-[14C]-U-phenylalanine
(n=8) and cycloheximide (n=8) were used to quantify protein synthesis and
degradation, respectively. Western blotting analyses was performed to determine
the activation of protein synthesis and degradation pathways. During hyperosmotic stress, protein degradation increased (p<0.05), while protein synthesis
was decreased (p<0.05) as compared to the iso-osmotic condition. The decline in
protein synthesis was accompanied by a decrease (p<0.05) in p70s6 kinase
phosphorylation, while the increase in protein degradation was associated with
an increase (p<0.05) in autolyzed calpain. Therefore, hyper-osmotic extracellular
stress results in an intracellular catabolic environment in mammalian skeletal
muscle tissue.
i TABLE OF CONTENTS ABSTRACT ................................................................................................................ i TABLE OF CONTENTS ........................................................................................... ii LIST OF FIGURES .................................................................................................... v LIST OF TABLES .................................................................................................... vi LIST OF ABBREVIATIONS ................................................................................... vii CHAPTER 1.1: WATER – THE CELLULAR BASIS OF LIFE ............................... 1 I. Introduction ...................................................................................................................... 1 II. Compartmentalization of Fluids ..................................................................................... 1 III. Osmolality ...................................................................................................................... 2 VI. Water Transport ............................................................................................................ 4 V. Whole Body Hydration and Fluid Shifts ........................................................................ 6 CHAPTER 1.2: CELL VOLUME DETECTION AND REGULATION ................... 10 I. Cell Volume Detection .................................................................................................. 10 Ia. Mechanical Cell Volume Detection ............................................................................. 10 Ib. Osmotic Cell Volume Detection .................................................................................. 11 II. Cell Volume Regulation ............................................................................................... 12 IIa. Regulatory Volume Increase ...................................................................................... 12 III. Cell Volume and Metabolism ...................................................................................... 14 CHAPTER 1.3: SKELETAL MUSCLE AND PROTEIN METABOLISM .............. 16 I. Fluid Shifts in Skeletal Muscle ...................................................................................... 17 II. Cell Volume and Skeletal Muscle Metabolism ........................................................... 19 III. Protein Degradation Pathways ................................................................................... 21 ii IIIa. Ubiquitin Proteasome System ................................................................................... 21 IIIb. The Calpain System ................................................................................................... 25 IV. Protein Synthesis ........................................................................................................ 29 V. Summary ...................................................................................................................... 34 CHAPTER 2: STATEMENT OF THE PROBLEM ................................................. 36 I. Statement of the Problem ............................................................................................. 36 II. Purpose ........................................................................................................................ 36 III. Hypotheses ................................................................................................................. 37 CHAPTER 3: METHODOLOGY ............................................................................. 38 I. Animals and Housing .................................................................................................... 38 II. Osmotic Conditions ...................................................................................................... 39 III. Muscle Extraction ........................................................................................................ 39 IV. Protein Turn-over Measures ...................................................................................... 41 V. Western Blotting Analysis: .......................................................................................... 43 VI. Metabolite Analysis: .................................................................................................... 45 VII. Statistical Analysis ..................................................................................................... 46 CHAPTER 4: RESULTS ......................................................................................... 47 I. General Observations ................................................................................................... 47 II. Relative Water Content ................................................................................................ 47 III. Metabolites .................................................................................................................. 48 IV. Protein Synthesis ........................................................................................................ 48 V. Protein Degradation ..................................................................................................... 49 VI. Western Blotting Analyses ......................................................................................... 50 CHAPTER 5: DISCUSSION ................................................................................... 55 iii I. General Observations ................................................................................................... 56 II. Protein Synthesis ......................................................................................................... 58 III. Protein Degradation .................................................................................................... 62 IV. Summary and Conclusions ........................................................................................ 66 V. Future Directions ......................................................................................................... 69 REFERENCES: ....................................................................................................... 71 APPENDIX I: BREAKDOWN OF MODIFIED SIGMA MEDIA-199 ...................... 86 APPENDIX II: LABORATORY PROCEDURES ................................................... 88 iv LIST OF FIGURES FIGURE 1. The discovery of aquaporins in Xenopus laevis oocytes – p. 6
FIGURE 2. Water loss from bodily compartments – p. 9
FIGURE 3. Cell shrinkage and protein turnover in hepatocytes – p. 16
FIGURE 4. Protein synthesis during hyper-osmotic stress in muscle cell culture
– p. 21
FIGURE 5. The ubiquitin proteasome system – p. 23
FIGURE 6. Hypothesized findings of the current study – p. 34
FIGURE 7. Experimental design – p. 38
FIGURE 8. Skeletal muscle incubation bath system – p. 41
FIGURE 9. Relative muscle water weight – p. 48
FIGURE 10. Incorporated phenylalanine during hyper-osmotic stress – p. 49
FIGURE 11. Tyrosine release during hyper-osmotic stress – p. 49
FIGURE 12. Total mTOR protein content during hyper-osmotic stress – p. 51
FIGURE 13. Ser 2448 mTOR phosphorylation during hyper-osmotic stress – p.
51
FIGURE 14. Total p70s6 kinase protein content during hyper-osmotic stress – p.
52
FIGURE 15. Thr 389 p70s6 kinase phosphorylation during hyper-osmotic stress
– p. 52
FIGURE 16. Total actin protein content during hyper-osmotic stress – p. 53
FIGURE 17. Total ubiquitinated protein during hyper-osmotic stress – p. 53
FIGURE 18. Calpain autolysis during hyper-osmotic stress – p. 54
FIGURE 19. Summary of protein turnover findings during hyper-osmotic stress –
p. 69
v LIST OF TABLES
TABLE 1. Rodent Weights and Media Osmolality – p. 47
TABLE 2. High Energy Phosphate Concentrations – p. 48
vi LIST OF ABBREVIATIONS
ADH – Antidiuretic Hormone
MAP – Mitogen Activated Protein
ANP – Atrial Natriuretic Peptide
Na+ – Sodium
APS – Ammonium Persulfate
NaCl – Sodium Chloride
AQP – Aquaporin Water Channel
Na2HPO4 – Sodium Phosphate
ATP – Adenosine Triphosphate
NaOH – Sodium Hydroxide
Br- – Bromide
NHE – Sodium/Hydrogen Exchanger
BSA – Bovine Serum Albumin
NKCC – Na+-K--2Cl- Cotransporter
Ca2+ - Calcium
NO3- – Nitrate
Cl- – Chloride
O2 – Oxygen
CO2 – Carbon Dioxide
Tween
PBST – Phosphate Buffered Saline
DHPR –Dihydropyridine Receptor
PC – Phosphatidylcholine
EDL– Extensor Digitorum Longus
PCr – Phosphocreatine
EGTA – Ethylene Glycol Tetraacetic
Acid
PI - Phosphatidylinositol
GLUT – Glucose Transporter
PI3K – Phosphatidylinositol-3-kinase
H+ – Hydrogen Ion
PVDF – Polyvinylidene Fluoride
HYPER – Hyper-osmotic
RBC – Red Blood Cell
I- – Iodide
RVD – Regulatory Volume Decrease
IGF-1 – Insulin-like Growth Factor
RVI – Regulatory Volume Increase
ISO – Iso-osmotic
SDS – Sodium Dodecyl Sulfate
K+ – Potassium
SR – Sarcoplasmic Reticulum
KCC – K+-Cl- Cotransporter
TBST – Tris Buffered Saline Tween
KCl – Potassium Chloride
TEMED - Tetramethylethylenediamine
K2HPO4 – Potassium Phosphate
Channel
VRAC – Volume Regulated Anion
mTOR – Mammalian Target of Rapamyacin
vii CHAPTER 1.1: WATER – THE CELLULAR BASIS OF LIFE
I. Introduction
Water is the one of the most important molecules in the human body,
where it accounts for approximately 55-60% of total body weight (Macknight and
Leaf 1977). It is vital for a variety of functions that are critical to survival such as
the regulation of pH and body temperature. Water is also involved in the delivery
of O2/nutrients to cells throughout the body, as it is a large component of blood.
Blood plasma consists of approximately 90% water, with the other 10% made up
of dissolved proteins, clotting factors, glucose, hormones, mineral ions and CO2.
Another key function of water is the ability to act as a solvent. Sugars, salts,
amino acids, minerals and an array of other nutrients/molecules are dissolved in
water, which provides an optimal medium for cellular reactions to occur. The
solvent properties of water can also lead to trans-membrane fluid shifts between
tissues and cells of the body (See Osmolality section). As noted by the examples
above, it is clear that adequate hydration is necessary for the maintenance of
homeostasis and optimal bodily function.
II. Compartmentalization of Fluids
Water is not evenly distributed throughout the body, as it is separated into
intracellular and extracellular compartments. Intracellular water is comprised of
all water that is located within the cells of the body and accounts for 2/3 of total
body water. It provides the base for the cytoplasm, which in turn, provides the
medium for metabolic reactions and other cellular processes to occur.
Extracellular water is all water that is not located within the cells of the body. It
accounts for the remaining 1/3 of total body water. The extracellular compartment
can be further sub-divided into the plasma, interstitial and lymphatic
compartments. Plasma is the liquid component of blood and consists of
approximately 90% water. Interstitial fluid is the fluid that surrounds the cells of
the body. It provides a means for delivering nutrients from the bloodstream to the
cell as well as removing any waste that is released by the cell and returning it to
the vasculature. Interstitial fluid may also have a protective role for certain cells in
the body (e.g. brain cells) as the fluid cushions the cell during forceful impacts.
The lymphatic system makes up the final part of the extracellular compartment,
and is defined as a collection of ducts, tissues, and nodes that are spread
throughout the body and carry lymph within them. Lymph is similar to blood
plasma in composition but also contains white blood cells. While the lymphatic
system has multiple functions, its main function in terms of water balance is to
return excess interstitial fluid to the blood (Swartz and Boardman 2002). Excess
interstitial fluid can lead to trans-membrane fluid shifts and subsequent cell
volume changes, which can affect cellular processes therefore the lymphatic
system is also vital for water balance.
III. Osmolality
Osmolality and osmolarity are two terms that are often used
interchangeably but are unique measures. Osmolarity is the amount of dissolved
solute in a solution. Since it is relative to the amount of solution, it is measured in
osmoles per litre (Osm/L). Osmolality, on the other hand, is the amount of solute
2 dissolved per kilogram of solvent (Osm/kg). The volume of a solution will change
with the addition of solutes or with changes in temperature and pressure, thereby
influencing osmolarity. The weight of solvent remains the same regardless of
temperature, pressure, or solutes added; therefore osmolality is a more constant
measure.
In the body, normal serum osmolality varies between 285-295 mmol/kg of
water, but environmental stressors (e.g. exercise, diet, hydration, diabetes) can
lead to altered extracellular osmolalities which subsequently lead to trans-cellular
fluid shifts (Kraft 2000). There are three different physiological extracellular
osmotic states that can influence intracellular volume. A hypotonic extracellular
osmotic environment (<285mmol/kg) occurs when there are a fewer number of
solutes in the extracellular space than are located within the cytoplasm. To
equilibrate this osmotic gradient, water is rushed into the cell resulting in a net
increase in cell volume. A hypertonic extracellular osmotic environment
(>295mmol/kg) occurs when there are a greater number of solutes in the
extracellular media than within the cytoplasm. Ultimately, water leaves the cell
and cell volume decreases. Finally, an iso-osmotic extracellular environment
occurs when there are an equal amount of solutes in the cytoplasm and
extracellular space therefore an equal amount of water enters and exits the cell
leaving cell volume constant. This is the favoured osmotic state of the cell. It
appears that cell volume is well controlled, and that cells possess homeostatic
mechanisms which attempt to maintain cell volume at a preferred cell size
(Okada 2004) (See Cell Volume Regulation Section).
3 VI. Water Transport It was initially believed that water moved between body compartments
only by way of simple diffusion. Extensive research on an assortment of tissues
has established that different tissues have varying water permeabilities (Nielsen,
Chou et al. 1995; Koyama, Yamamoto et al. 1997). These findings lead
researchers to investigate the existence of other possible water pathways. It is
now known that there are three routes of trans-membrane water movement:
simple diffusion, co-transport, and aquaporins (Borgnia, Nielsen et al. 1999).
Simple diffusion is the passing of a substance from an area of high
concentration to an area of low concentration through a membrane without the
aid of a membrane protein. Water passes through the hydrophobic phospholipid
bi-layer but is highly dependent on the presence of a large concentration
gradient. Since it is a passive process and the membrane is partially
hydrophobic, the process is relatively slow.
The second method of water movement is that of co-transport. This occurs
when water is moved across a cell membrane with other molecules. Different
molecules and ions possess varying osmotic effects; therefore water will tend to
follow certain ions and molecules more than others as they are passed through a
membrane. An example of this involves the NKCC co-transporter. This symporter
is responsible for moving Na+, K+, and two Cl- ions across the membrane in
addition to water (Gosmanov, Lindinger et al. 2003). Co-transport is an effective
method for mass water transport but is dependent on other cellular processes.
These symporters are not specific to water therefore co-transport is dependent
4 on the ion concentrations inside and outside of the cell. It is the ion flux which is
regulated in this instance, not the water concentration.
The final pathway for water transport is through aquaporin channels.
Aquaporins were first discovered in the late 1980’s by accident when researchers
were attempting to identify Rh blood group antigens (Agre, Saboori et al. 1987).
The protein fragment was analyzed and purified over the next few years and
eventually determined to have channel-like properties that were specific to water
(Smith and Agre 1991). Researchers confirmed these findings using oocytes
expressing the newly found protein. The oocytes were placed into a hypotonic
solution where they swelled until exploding. It was later found that their
coefficient of osmotic water permeability was 20 times that of control oocytes
(See Figure 1) (Preston, Carroll et al. 1992). This marked the discovery of the
water channel and the term aquaporin was coined (Agre, Sasaki et al. 1993).
Aquaporins have since been found in plants, animals and microorganisms. To
date, there have been 13 different aquaporin proteins identified in the human
body. The two main classifications of in the body are: aquaporin and
aquaglyceroporin. Aquaporin 1, 2, 4, 5, and 8 only allow water to flow through
them, while aquaglyceroporin 3, 7, 9, and 10 move water in addition to other
small solutes such as glycerol and urea (Verkman 2005). It is estimated that 3 X
109 water molecules pass through a given aquaporin channel in one second,
which makes them the primary source of water transport in the body (Zeidel,
Ambudkar et al. 1992). In humans, these water channels can be found in a
variety of tissues including: brain, skeletal muscle (which has primarily AQP-4),
5 liver, but most notably in the kidney. Aquaporin 1, 2, 3, 4, 6, 7, and 8 can all be
found in various parts of the kidney, primarily within the nephron. While
aquaporin throughout the body vary slightly in terms of function, they work
collectively to maintain body water balance (Nielsen, Frokiaer et al. 2002).
FIGURE 1. The discovery of aquaporins in Xenopus laevis oocytes. Cells were
injected with AQP1 RNA and placed in a hypo-osmotic environment where cell
swelling occurred (Preston, Carroll et al. 1992)
V. Whole Body Hydration and Fluid Shifts Whole body water balance is largely regulated through the endocrine
system. The main endocrine hormones involved include: anti-diuretic hormone
(ADH), angiotensin II and atrial natriuretic peptide (ANP). ADH is a peptide
hormone that is released from the posterior pituitary gland and acts primarily on
the kidneys. Upon an increase in plasma osmolality, osmoreceptors in the
hypothalamus signal to the pituitary gland to release ADH into the bloodstream.
Once reaching the kidneys, ADH increases the permeability of the collecting duct
in the nephron to allow increased reabsorption of water (Bonjour and Malvin
1970). This action helps to restore iso-osmotic conditions in the plasma. During
6 conditions of hypo-osmotic plasma, ADH secretion by the posterior pituitary
gland is suppressed.
Angiotensin II and ANP also play a large role in whole body water
regulation. When blood volume is low (e.g. dehydration), the kidney secretes
renin into circulation. Renin will travel in the bloodstream until it reaches the liver
where it then facilitates the conversion of angiotensinogen into angiotensin I.
Angiotensin I is released by the liver into circulation and is subsequently
converted into angiotensin II by angiotensin converting enzyme, which can be
found in the lungs. Upon its activation, angiotensin II acts on blood vessels,
causing vasoconstriction, and thereby increasing blood pressure. In terms of
water balance, angiotensin II also stimulates an increase in the release of the
hormone aldosterone. Aldosterone causes Na+ ions and H2O to be reabsorbed
from the distal convoluted tubule and collecting duct of the nephron into
circulation. This action helps to restore whole body hydration levels. ANP is also
involved in whole body water balance. Once iso-osmotic conditions are achieved
in plasma, ANP is released into circulation by the heart. ANP acts to suppress
the release of renin and aldosterone (Antunes-Rodrigues, de Castro et al. 2004).
Together, ADH, angiotensin II and ANP work to regulate water intake and
excretion.
Fluid shifts between the plasma and interstitium are affected by two
forces: hydrostatic and colloid osmotic forces. Hydrostatic pressure is the
pressure generated by fluid pushing against the walls of the blood vessels (Reed
1981). This results in fluid being forced out of the blood vessels and into the
7 interstitium. Colloid osmotic pressure is the force that draws fluid from the
interstitium back into the vasculature. It is primarily due to the osmotic effect of
blood proteins. These proteins are too large to leave the capillaries but possess
an osmotic effect that helps to draw water into the capillaries. Together,
hydrostatic pressure and colloid osmotic pressure provide a balance of forces
that equilibrates fluid between the capillaries and the interstitium.
Similar to the rest of the body, fluid shifts are constantly occurring between
intracellular and extracellular water compartments. Generally, variations in
extracellular osmotic environment are responsible for trans-membrane fluid
shifts. These environments can be the result of a variety of stimuli such as
physical exercise, diet, hormones and injury/disease. In conditions of cell
shrinkage (e.g. exercise, dehydration) it is suggested that fluid shifts serve to
protect cardiovascular function first and foremost, as maintenance of blood
volume is critical for regulation of arterial blood pressure and thermoregulation
(Fortney, Nadel et al. 1981). During exercise, blood volume is initially decreased
as fluid is released through pores of the skin as sweat. Although sweat
production provides a thermoregulatory role for the body, cardiac output and
stroke volume are decreased which diminishes endurance exercise capacity
(Fortney, Wenger et al. 1983). Optimal blood transport cannot be sustained with
these declines in plasma therefore blood volume must be replenished. In the
long-term, water becomes redistributed within its compartments to return blood
plasma to an adequate fluid level. This restoration of plasma occurs with or
without fluid replacement as the increasing osmolality of blood plasma (due to
8 fluid loss from sweat and increased concentration of Na+) creates an osmotic
gradient that draws water to the vasculature (Sanders, Noakes et al. 2001).
Water follows an osmotic gradient from the intracellular compartment to the
interstitium to the blood plasma. As fluid loss worsens, there is a
disproportionately greater water loss from the intracellular compartment in order
to maintain blood volume (See Figure 2) (Costill, Cote et al. 1976; Durkot,
Martinez et al. 1986)
11% 4 Total Body Water Loss (L) 3.5 10% 3 2.5 39% 38% 2 Plasma 10% 1.5 50% 60% Intracellular 52% 1 Inters��al 30% 0.5 0 -­‐2.2 30% -­‐4.1 -­‐5.8 Level of Dehydra�on (%) FIGURE 2. Water loss from bodily compartments during dehydration. As the
body dehydrates, total body water lost is increased. As dehydration worsens, a
higher % of water is lost from the intracellular compartment. Adapted from
Costille, Cote et al. 1976; Durkot, Martinez et al. 1986.
9 CHAPTER 1.2: CELL VOLUME DETECTION AND REGULATION
I. Cell Volume Detection
There is no consensus as to how cells detect changes in cell volume.
However, two different theories have been proposed: mechanically-induced and
osmotically-induced cell volume detection.
Ia. Mechanical Cell Volume Detection Mechanically induced cell volume detection has been suggested to occur
as the result of deformation of the cell membrane due to extracellular osmotic
stress (Parker 1993). In this theory, the cell membrane experiences either
stretching (cell swelling) or bending (cell shrinkage) and results in a rearrangement of the cytoskeleton within the cell. These perturbations result in
intracellular signals being transmitted through the cell which activate
mechanisms to attempt to return the cell to its preferred cell volume. In terms of
water transport specifically, it is hypothesized that aquaporin and ion channels
are recruited to the membrane (similar to GLUT-4 translocation) in order to move
water (Sachs 1991).
Research in red blood cells (RBC’s) has generated uncertainty around the
theory of mechanical cell volume detection. RBC’s have been shown to swell to
1.6 times their normal cell volume without experiencing any cell membrane
stretch. This is attributed to their biconcave shape. However, KCl channels,
which are known to be a main mechanism of water efflux following cell swelling,
were shown to become activated well before RBC’s experienced membrane
stretch (Jennings and Schulz 1990). These findings contradict the theory of
10 mechanical cell volume detection. The theory was further tested by researchers
using ghost cells (resealed RBC’s with the same cytoplasmic protein
concentration and surface area but a smaller cell volume) and control RBC’s.
The researchers compared the two cell types in regulatory volume changes (KCl
activation) and observed that both cell types underwent KCl channel activation
upon changes in intracellular osmolality, whereas the difference in cell volume
did not have an effect (Colclasure and Parker 1992). Although these findings
oppose the theory of mechanical cell volume detection, results from RBC’s may
not translate to other cell types, therefore further research is warranted.
Ib. Osmotic Cell Volume Detection Osmotically induced cell volume detection occurs when impermeable
components of the cell (i.e. proteins, carbohydrates, lipids, and nucleic acids)
change in concentration due to the alterations in fluid content, which is then
detected by the cell (Sarkadi and Parker 1991). It is hypothesized that these
concentration changes will trigger intracellular signalling pathways and
mechanisms to return the cell to its normal cell volume (Sarkadi and Parker
1991). This theory is in accordance with the theory of macromolecular crowding;
the crowding that occurs within the cell due to the large presence of
macromolecules within the cytoplasm. Macromolecular crowding has been
shown to affect the properties of macromolecules and enzymes which
subsequently influence chemical processes (e.g. protein transcription) within the
cell (Zimmerman 1993; Garner and Burg 1994).
11 II. Cell Volume Regulation Many systems in the human body are homeostatically regulated and cell
volume is no different. It has been established that cell volume is dynamic but in
order to maintain optimal function, the cell will attempt to maintain a stable
volume. In addition, evidence suggests that cell volume can act as a signal within
the cell (see Cell Volume and Metabolism section) therefore cell volume must be
tightly controlled. To achieve this, there are mechanisms in place which attempt
to return the cell to its original cell volume. These general mechanisms are
known as Regulatory Volume Decrease (RVD) and Regulatory Volume Increase
(RVI). When a cell is placed in a hypo-osmotic solution the cell will initially swell
and RVD mechanisms are then engaged. When a cell is placed in a hyperosmotic solution the cell will initially shrink followed by activation of RVI
processes. Both mechanisms are acute and represent rapid attempts to return
cell volume to a homeostatic level. Since cell shrinkage is the focus of the current
study, I will concentrate solely on RVI mechanisms.
IIa. Regulatory Volume Increase No single mechanism is responsible for water influx into the cell following
cell shrinkage but instead a variety of channels, transporters and molecules are
used. The major mechanisms include: the Na+/H+ exchange port (NHE), Na+-K+2 Cl- co-transporter (NKCC1) and organic osmolytes. Upon cell shrinkage, it is
speculated that NHE’s are activated to allow Na+ to enter the cell while H+ is
extruded. The proposed mechanism is a decrease in cell volume will increase the
affinity of an allosteric site on NHE for intracellular H+, which in turn will activate
12 the exchanger (Grinstein, Rothstein et al. 1985). The NKCC is similarly reliant on
Na+ to initiate water influx. This transporter is responsible for transporting one
Na+, one K+, and two Cl- ions into the cell. This action helps to increase
intracellular osmolality and promote the flow of water into the cell (Haas and
Forbush 1998). The activation of this transporter is believed to be due to
phosphorylation of the transporter during hyper-osmotic induced cell shrinkage
(Gagnon, England et al. 2007).
During RVI, elevated concentrations of intracellular ions have the potential
to negatively influence cell function (e.g. fluctuation of pH, function of proteins,
denaturation of macromolecules, alteration of resting membrane potential)
(McManus, Churchwell et al. 1995). Organic osmolytes provide an alternative
route of cell volume maintenance that is less stressful to the cell as ion flux is not
involved. Osmolytes are organic compounds that can either be taken up by or
synthesized within the cell and function to increase intracellular osmolality. As
osmolytes accumulate, intracellular ions can be released without sacrificing cell
volume. There are three main classes of organic osmolytes: polyols (sorbitol,
myoinositol and inositol), methylamines (betaine, glycerophosphorylcholine), and
amino acids (taurine, glutamine, glutamate, glycine and aspartate) (Lang, Busch
et al. 1998). Different cell types make use of different osmolytes. Due to the
adverse long-term effects of ion entry, ion flux is generally seen as an acute
mechanism of RVI, whereas organic osmolyte accumulation provides a more
long-term solution to cell volume (Lang, Busch et al. 1998).
13 Cell volume regulation is an extremely complex process with multiple
mechanisms involved. It is important to note that different cell types will regulate
cell volume differently. In order to maintain optimal function, cell volume must be
tightly controlled as cell volume fluctuations have been shown to influence
intracellular reactions within the cell, particularly metabolic reactions (see Cell
Volume and Metabolism section).
III. Cell Volume and Metabolism Extensive research on hepatocytes in the early 1990’s has shown that
cellular hydration influences metabolic pathways within the cell (Haussinger,
Lang et al. 1994). This research led to the development of the cell swelling
theory, which states that cellular volume acts as a signal and has control over
metabolic pathways within the cell. More specifically, cellular swelling creates an
anabolic environment, while cellular shrinkage promotes an intracellular catabolic
environment. The reason for these metabolic alterations are unknown but have
been hypothesized to serve as a form of osmoregulation. During cell swelling, the
synthesis of macromolecules reduces the osmotic effect of smaller organic
molecules thereby limiting water influx. The opposite is true for cell shrinkage; the
degradation of macromolecules into smaller organic molecules increases the
overall osmotic effect and promotes the uptake of water. The remainder of this
thesis will focus on cell shrinkage.
Metabolic pathways and intracellular metabolites (ATP, PCr) have been
shown to be affected by hyper-osmotic induced cellular shrinkage in a variety of
tissues (Haussinger 1996; Antolic, Harrison et al. 2007). Within carbohydrate
14 metabolism, cell shrinkage has been shown to inhibit glycogen synthesis as well
as stimulate glycogen breakdown (Graf, Haddad et al. 1988; Baquet, Hue et al.
1990). There has been little research dedicated to hyper-osmotic induced cell
shrinkage and fat metabolism. However, protein turnover and hyper-osmotic
stress has received a great deal of attention in the academic community. In
hepatocytes, hyper-osmotic induced cell shrinkage has been shown to suppress
protein synthesis while stimulating protein degradation (See Figure 3) (Stoll,
Gerok et al. 1992). Researchers have confirmed these findings on a whole body
level (Berneis, Ninnis et al. 1999).
Cellular hydration has been shown to influence metabolic pathways and
processes within the cell as noted by the examples above. Of particular interest
is the suppression of protein synthesis and stimulation of protein degradation
during cell shrinkage. The majority of research in the area has been conducted
using hepatocyte models but applications in skeletal muscle could be significant
due to its large source of functional protein.
15 FIGURE 3. Cell shrinkage and protein turnover in hepatocytes. Cell shrinkage
creates an intracellular catabolic environment and negative net protein balance
(Stoll, Gerok et al. 1992)
CHAPTER 1.3: SKELETAL MUSCLE AND PROTEIN METABOLISM
The ability to contract and produce force makes skeletal muscle cells one
of the more unique cell types within the body. To support this property, skeletal
muscle contains a large store of myofibrillar protein. The two major proteins in
muscle, actin and myosin, interact to develop force through the cycling of crossbridges. Several other proteins serve to stabilize the structure of the sarcomere
(e.g. desmin, titin). Intracellular proteins are continually synthesized and
degraded in order to maintain optimal muscle function. To support this
regeneration, skeletal muscle contains the largest store of amino acids in the
body (Bergstrom, Furst et al. 1974). Factors such as muscular damage, amino
acid availability, disuse, cachexia, and disease have the ability to influence rates
of protein synthesis and degradation (Rennie 1985; Garibotto, Russo et al. 1994;
16 Cooney, Kimball et al. 1997; Phillips, Glover et al. 2009). Overall, skeletal muscle
contributes approximately 25–30% of whole body protein turnover (Balagopal,
Rooyackers et al. 1997).
Protein turnover involves two major mechanisms: protein synthesis and
protein degradation. Protein synthesis is the building of proteins and can be
further broken down into transcription and translation processes. Transcription is
the process of creating a complementary RNA copy from a specific DNA
sequence. Translation involves the recruitment of specific amino acids that
correspond to the transcribed mRNA molecule, followed by construction of the
protein at the level of the ribosomes. Conversely, protein degradation is the
digestion of proteins into amino acids. Proteins can be used as a nutritional
source but often it is older or damaged proteins that are targeted. Several
mechanisms can be used within muscle to degrade protein including: the
ubiquitin proteasome system and calpain system (See Protein Degradation
Section).
Skeletal muscle houses a large store of both functional protein and amino
acids. In addition, protein turnover in skeletal muscle is dynamic and can be
influenced by a variety of factors. For these reasons, skeletal muscle is an
attractive tissue to investigate in terms of protein turnover.
I. Fluid Shifts in Skeletal Muscle Although a number of factors can influence cellular hydration in skeletal
muscle, physical exercise is one of the main triggers. During exercise, significant
fluid shifts are seen between the vasculature, interstitium and intracellular
17 (muscle cell) compartments. In order to supply the working muscles with O2 and
nutrients, an increase in cardiac output is required, which in turn, increases blood
pressure. This results in an increase in hydrostatic pressure, which consequently,
forces fluid out of the capillaries and into the interstitial space (El-Sayed 1998). In
addition to these events, increased energy turnover within the muscle leads to a
build-up of metabolic by-products, thereby increasing intracellular osmolality
(Lindinger, Spriet et al. 1994). Altogether, these actions result in the net
movement of water from the capillaries into the active muscle cells, resulting in
an acute increase in muscle cell volume. However, during prolonged exercise,
plasma volume is decreased due to sweating. This results in a net decrease of
both total body fluid and muscle cell volume. Similar to other cell types, there is a
disproportionately greater water loss from the intracellular compartment in order
to maintain blood volume during prolonged exercise (See Figure 2) (Costill, Cote
et al. 1976; Durkot, Martinez et al. 1986).
Aside from exercise, other conditions (e.g. diabetes) can lead to skeletal
muscle fluid shifts. The signature characteristic of diabetes is hyperglycaemia
due to insulin unresponsiveness or deficiency. The chronic elevation of blood
glucose can have adverse effects on organs throughout the body including the
eyes, kidneys, and blood vessels. To counteract, the body will attempt to reduce
plasma blood glucose by excreting glucose through urine. As a result, total body
water is reduced and dehydration can occur. In addition, long-term diabetes
(Type I and II) has been shown to result in increased vascular permeability to
both small and large proteins in skeletal muscle (Trap-Jensen 1970; Parving and
18 Rossing 1973; Leinonen, Matikainen et al. 1982). As a result, proteins move from
the vasculature to the interstitium, thereby increasing interstitial osmolality. To
equilibrate, fluid is drawn from within the muscle cell resulting in muscle cell
shrinkage (Fauchald, Norseth et al. 1985).
II. Cell Volume and Skeletal Muscle Metabolism According to the theory of cell swelling, cell volume acts as a signal and
has influence over metabolic pathways within the cell. Over the past decade,
research has begun to elucidate this theory in skeletal muscle. Through an
isolated muscle model, researchers have demonstrated the effect of extracellular
osmolality on skeletal muscle cell volume as well as the responsiveness of
muscle fiber types to fluid shifts (Antolic, Harrison et al. 2007). Furthermore,
researchers have investigated the influence of extracellular hyper-osmotic stress
on glucose uptake as well as recovery of glucose kinetics from hyper-osmotic
stress. Glucose uptake was found to increase in isolated intact EDL muscles
after they being exposed to extracellular hyper-osmotic stress for 60 minutes; this
increase could be reversed by exposing the same muscle to iso-osmotic
extracellular media for 5 minutes (Farlinger 2007; Mulligan 2008). Despite
evidence of skeletal muscle as a large storage site for amino acids and functional
protein, as well as a tissue which faces osmotic challenges in a variety of
conditions (e.g. exercise and diabetes), little research has been dedicated to the
cell shrinkage and protein turnover in skeletal muscle, until recently when
researchers quantified total protein synthesis under osmotic stress (Roy 2009).
Using a cell culture model, L-6 muscle cells were placed in hyper-osmotic and
19 hypo-osmotic solutions and protein synthesis was subsequently measured. A
hyper-osmotic environment was found to suppress protein synthesis while no
changes were seen in a hypo-osmotic environment (See Figure 4). This
decrease in protein synthesis was attributed to the suspected catabolic state of
the cell. While this study was extremely novel in its findings, further research is
necessary to understand the bigger picture. Total protein degradation was not
measured in the experiment which results in an incomplete picture of protein
turnover. Secondly, although the cell culture model used provides a certain level
of control within an experiment, it is not very physiological. An isolated intact
skeletal muscle would provide a more physiological model as the structure of the
muscle is completely intact and viable. Lastly, measuring the activity of specific
protein anabolic and catabolic pathways would allow for a more in-depth
understanding of the effect of extracellular hyper-osmotic stress on protein
turnover in skeletal muscle.
20 FIGURE 4. Protein synthesis during hyper-osmotic stress in skeletal muscle cell
culture. L-6 muscle cells were placed under hypo-osmotic, hyper-osmotic or isoosmotic conditions for either 24 or 48 hours. Hyper-osmotic stress was shown to
suppress total protein synthesis at both time points. Hypo-osmotic conditions had
no effect on protein synthesis (Roy 2009).
III. Protein Degradation Pathways
IIIa. Ubiquitin Proteasome System
The ubiquitin proteasome pathway is one of the main systems of protein
degradation in skeletal muscle. It is a multi-unit complex known as the 26S
proteasome, which is comprised of three subunits: a core 20S unit and two 19S
subunits (Pickart and Cohen 2004). In this model, proteins become tagged for
degradation by linking ubiquitin to targeted proteins. The complex also consists
of three enzymes: E1, E2, and E3. There are three main steps in the ubiquitin
proteasome pathway; the first step involves E1, which is an ATP-dependent
enzyme that activates ubiquitin by forming a thiol ester bond between the E1
enzyme and ubiquitin (See Figure 5). This prepares the proteasome for protein
degradation as only proteins tagged with ubiquitin can be degraded. The second
21 step involves the transfer of activated ubiquitin from E1 to a specific cysteine
residue on E2 via another thiol ester bond. E2 also adds the first ubiquitin to the
targeted protein. There are several E2 enzymes in the proteasome complex that
can perform this function. The third step occurs when activated ubiquitin is
transferred from E2 enzymes to a cysteine residue located on the E3 enzyme
(Hershko and Ciechanover 1992). Once attached, E3 enzymes continue to add
multiple ubiquitin molecules to the targeted protein (poly-ubiquitination). The
protein cannot be degraded until at least four or more ubiquitin molecules have
been added the protein (Pickart 2000). At this point, the 26S proteasome will
degrade the targeted protein into polypeptides molecules. Ubiquitin molecules
are released from the poly-ubiquitin chain once the targeted protein has been
degraded and the ubiquitin molecules can be subsequently recycled and used in
the degradation of other proteins.
22 FIGURE 5. The ubiquitin proteasome system. Figure adapted from
http://www.cellsignal.com/reference/pathway/Ubiquitin_Proteasome.html. E1
enzyme activates ubiquitin by binding to it. E2 attaches the 1st ubiquitin to protein
of interest. E3 adds further ubiquitin molecules. 26S proteasome degrades
proteins into poly-peptide chains.
During skeletal muscle atrophy, the E1, E2 and E3 enzymes have been
shown to experience a change in expression, specifically E3 or ubiquitin ligases.
Several E3 ligases have been identified in a variety of cell types, but two appear
to be more important in skeletal muscle atrophy: MuRF1 and Atrogin-1. Using
immobilization and hind-limb suspension models in rodents, researchers found
an up-regulation of the Atrogin-1 gene in all cases. As a follow-up, an Atrogin-1
KO model was subjected to muscle denervation and it was found atrophy was
attenuated by 56% at 14 days (Bodine, Latres et al. 2001). In the same study,
MuRF1 was also identified as an E3 ligase which was up-regulated in all models
of skeletal muscle atrophy. These results were later confirmed using a muscle
unloading model (Stevenson, Giresi et al. 2003). The effects of these scientific
models, specifically hind-limb suspension, may be mediated by cellular
dehydration, due to the effects they have on skeletal muscle fluid balance.
23 Researchers have consistently shown that the hind-limb suspension model
results in a decrease of intracellular water content in skeletal muscle (Booth
1982; Aoki, Lima et al. 2004). This is partly due a reduction in muscle cell
diameter but with this comes a decrease in intracellular water. Additionally,
researchers have shown an increase of AQP 4 protein expression in skeletal
muscle during hind-limb suspension (Basco, Nicchia et al. 2010). This increase is
believed to be linked to the efflux of water from within the cell. Although hind-limb
suspension does not directly represent cellular shrinkage, the model does
represent a loss of intracellular water.
Another environment where muscle cell shrinkage and increased
protein degradation are present is space flight (micro-gravity). On Earth, most
body water is located in the lower extremities due to gravitational pull. Upon
entering micro-gravity conditions, water becomes redistributed within the body
resulting in a head-ward shift of fluid and a subsequent decrease in muscle cell
volume in the lower half of the body (Moore and Thornton 1987). In addition to
fluid redistribution, space flight has been shown to result in an up-regulation of
the ubiquitin proteasome system. Rodents were shuttled into space and exposed
to micro-gravity for 2 hours before euthanasia. Lower limb muscles were excised
and western and northern blotting analyses was performed. Researchers found
an increase in ubiquitin mRNA as well as ubiquitinated protein. In addition, RC2
and RC9, subunits of the proteasome, were found to have an increased
expression (Ikemoto, Nikawa et al. 2001). The cause of the up-regulation of the
24 proteasome remains unclear, but it is possible that skeletal muscle cell hydration
may be contributing to the regulation of this system.
The ubiquitin proteasome pathway is one of the major pathways for
protein degradation within skeletal muscle. Certain scientific models (e.g. hindlimb suspension, micro-gravity) which result in cell volume shrinkage have shown
an up-regulation of the system. Therefore, hyper-osmotic induced cell shrinkage
in skeletal muscle could potentially result in increased activation of the ubiquitin
proteasome system as well as an increase in total protein degradation.
IIIb. The Calpain System
The calpain system is another major protein degradation pathway found in
skeletal muscle (Dayton, Goll et al. 1976). Calpains are cysteine proteases which
are activated by the presence of Ca2+ (Guroff 1964). Three different calpains are
found in skeletal muscle: the ubiquitous µ-calpain and m-calpain, and the muscle
specific p94 (Goll, Thompson et al. 2003; Bartoli and Richard 2005). There has
been limited research conducted on p94 due to the technical difficulty in isolating
the isoform. p94 has also been shown to undergo rapid autocatalytic degradation
after protein translation which makes it a challenge to detect and quantify in
skeletal muscle (Sorimachi, Toyama-Sorimachi et al. 1993). In addition, there
remains debate in the literature about the role Ca2+ plays in its activation. Some
believe that p94 activation is Ca2+ dependent while others believe that it can be
activated in the absence of Ca2+. Further research on p94 is required.
Calpains have several functions within muscle including cell migration,
apoptosis, cell proliferation as well as myofibrillar protein degradation. They have
25 been speculated to be localized around the Z-disk region in skeletal muscle
(Kumamoto, Kleese et al. 1992). µ and m calpains are homologous heterodimers
composed of an 80kDa unit and 30kDa subunit. The larger subunit contains the
catalytic domain while the smaller subunit is responsible for regulatory functions
(Smith, Lecker et al. 2008). The major difference between the two isoforms is the
Ca2+ concentration required for activation. µ calpain requires approximately 3-50
µM free Ca2+ while m calpain requires approximately 400-800 µM free Ca2+ to
achieve ½ proteolytic activity (Pontremoli, Viotti et al. 1990; Melloni, Michetti et
al. 1998; Goll, Thompson et al. 2003). In skeletal muscle, resting cytosolic Ca2+
concentration hovers around 100 nM, with increases up to 3-15 µM during an
action potential (Williams, Head et al. 1990; Delbono and Stefani 1993). Since
physiological Ca2+ peak tetanus does not reach the threshold for m calpain
activation, µ calpain will be the focus for this study.
Although there has been extensive research on calpain activation in a
variety of cell types, the mechanism is still debated within the academic
community. The following mechanism has been most accepted by researchers.
As intracellular Ca2+ concentrations rise, calpain will translocate to the cell
membrane where µ calpain will either bind a) a cell membrane phospholipid or b)
a free Ca2+ molecule. Evidence has shown that phospholipids within the cell
membrane, specifically the acidic phosphatidylinositol (PI), phosphatidylserine
(PS) and phosphatidylcholine (PC), have the potential to enhance µ calpain
activation by lowering the Ca2+ threshold required for activation (Coolican and
Hathaway 1984; Pontremoli, Melloni et al. 1985; Chakrabarti, Dasgupta et al.
26 1990). The proposed involvement of phospholipids is not completely supported
as a physiological mechanism has yet to be established, therefore further
research is necessary. Once µ calpain has bound to either a membrane
phospholipid or a free Ca2+ molecule, µ calpain will undergo a conformational
change resulting in the dissociation of its 80kDa and 30kDa subunits (Suzuki,
Hata et al. 2004). Paired with this dissociation, is a self-digestion of the 80kDa
subunit to 78kDa and 76kDa isoforms (Goll, Thompson et al. 2003). This selfdigestion is known as autolysis and is often used as indicator of calpain activity
because of its close association with calpain activation. During autolysis, a new
Ca2+ binding site is formed on the calpain protease domain. Ca2+ will bind to this
newly formed site and activate µ calpain to degrade intracellular proteins (Suzuki,
Hata et al. 2004).
Calpastatin is the main inhibitor of calpain activity. Eight calpastatin
isoforms have been identified to date in varying tissues, with each containing four
inhibitory domains, capable of binding one calpain each (Imajoh, Kawasaki et al.
1987; Parr, Sensky et al. 2001). Although an inhibitor, calpastatin still requires
Ca2+ in order to bind to calpain (Cottin, Vidalenc et al. 1981). In fact, calpastatin
requires less Ca2+ to bind to calpain than is required for calpain activation. To
avoid extensive inhibition by calpastatin, the protein does not bind Ca2+ directly
but instead is reliant on the Ca2+ associated with the calpain (Kapprell and Goll
1989). This allows calpains to be activated as required by the cell.
Calpains have been found to initiate the degradation of several myofibrillar
proteins in vitro including tropomyosin, troponin I, troponin T, desmin, titin and
27 nebulin (Toyo-Oka, Shimizu et al. 1978; Huang and Forsberg 1998). Evidence
exists to show myosin and actin are not targets of µ calpain (Toyo-Oka and
Masaki 1979). It is believed that calpains’ role is in the initiation of myofibrillar
protein degradation which results in the disorganization of myofibrils. From here,
the ubiquitin-proteasome-system is proposed to take over and further degrade
myofibrillar proteins (Solomon and Goldberg 1996).
Before calpain activation can occur, an increase in intracellular Ca2+
concentration is required. However, resting skeletal muscle typically contains low
concentrations of myoplasmic intracellular Ca2+ (~50nM), which are not great
enough to promote calpain activity. Therefore, if calpain activity were to be
altered during hyper-osmotic induced cell shrinkage, another source of Ca2+ is
required. The phenomenon of Ca2+ sparks were first seen in cardiac muscle but
their presence in skeletal muscle has subsequently been discovered (Cheng,
Lederer et al. 1993; Tsugorka, Rios et al. 1995). Ca2+ sparks are brief and
localized increases in myoplasmic Ca2+ concentrations and occur in the absence
of an action potential. One of the known causes of spontaneous Ca2+ sparks is
an extracellular hyper-osmotic environment. Researchers have found that
isolated and intact skeletal muscle fibers which were exposed to a hyper-osmotic
solution exhibited Ca2+ sparks (Martin, Petousi et al. 2003; Wang, Weisleder et
al. 2005). Researchers have proposed that hyper-osmotic induced cell shrinkage
results in an alteration of the structural interaction between DHPR and ryanodine
receptors resulting in the spontaneous release of Ca2+ from the SR (Apostol,
Ursu et al. 2009). In support of their hypothesis, the inhibition of DHPR activation
28 resulted in the suppression of Ca2+ sparks, which suggests that the Ca2+ sparks
are being released from intracellular sources (Pickering, White et al. 2009).
The role of calpains within skeletal muscle protein degradation has been
examined extensively in the past and evidence exists to show that they do play a
role in the degradation of sarcomeric proteins (Huang and Forsberg 1998; Goll,
Thompson et al. 2003). The phenomenon of Ca2+ sparks in skeletal muscle
during hyper-osmotic stress makes calpains an obvious candidate for upregulation in hyper-osmotic stress.
IV. Protein Synthesis
Protein synthesis is the process of building new proteins to replace old or
dysfunctional proteins. Before transcription and translation can occur, signalling
cascades must be initiated to stimulate the synthesis of specific proteins which
are required by the cell. One of the primary protein synthesis signalling pathways
in skeletal muscle is the Akt/mTOR/p70s6k pathway. Increased activity of this
pathway has been shown to result in hypertrophy while decreased flux through
the pathway leads to skeletal muscle atrophy (Bodine, Stitt et al. 2001).
At the whole body level, the regulation of protein synthesis begins with
amino acid availability as they are the building blocks of all proteins (Millward,
Nnanyelugo et al. 1974). However, certain hormones and growth factors (e.g.
IGF-1 and insulin), also have influence on activity of the Akt/mTOR/p70s6k
pathway. Upon the binding of IGF-1 and insulin to their respective receptors on
the cell membrane, phosphatidylinositol-3-kinase (PI3K) is activated. Once PI3K
is activated, it then will phosphorylate a membrane phospholipid,
29 phosphatidylinositol-4, 5-bisphosphate, to generate phosphotidylinositol-3, 4, 5trisphosphate; this action results in the formation of a binding site on
phosphatidylinositol-3, 4, 5-trisphosphate for Akt (Vivanco and Sawyers 2002;
Matsui, Nagoshi et al. 2003). Akt is a major serine/threonine kinase that is
involved in skeletal muscle protein synthesis through several routes of action. For
Akt activation to occur, Akt must translocate to the cell membrane and bind to
phosphotidylinositol-3, 4, 5-trisphosphate which results in its phosphorylation
(Stitt, Drujan et al. 2004). During models of skeletal muscle hypertrophy, the
proportion of phosphorylated Akt has been shown to be increased. Conversely,
rodent hind-limb suspension models have shown decreases in Akt
phosphorylation (Lai, Gonzalez et al. 2004).
Downstream from Akt, there are three main targets, all of which influence
protein synthesis. GSK-3-β (glycogen synthase kinase-3-beta) is responsible for
the synthesis of glycogen within skeletal muscle, but also blocks protein
translation when in its active form (Rommel, Bodine et al. 2001). Phosphorylated
Akt has been shown to inhibit the activation of GSK-3-β in myotubes (Cross,
Alessi et al. 1995). FOXO (Forkhead Box Proteins) are another downstream
target of Akt. FOXO proteins are transcription factors in skeletal muscle that
translocate to the nucleus upon activation and act to influence gene expression.
FOXO proteins appear to aid in the up-regulation of E3 ligases, MuRF-1 and
Atrogin-1 (Sandri, Sandri et al. 2004). As previously mentioned, MuRF-1 and
Atrogin-1 are involved in the process of poly-ubiquitination in skeletal muscle.
Activated Akt has been shown to inhibit the translocation of FOXO proteins to the
30 nucleus thereby negating their effect on the E3 ligases. Conversely, when Akt is
inactive, FOXO proteins are free to translocate to the nucleus and increase E3
ligase expression (Leger, Cartoni et al. 2006). The final and most important
downstream target of Akt is mammalian target of rapamyacin (mTOR). mTOR is
another serine/threonine kinase that is heavily involved in protein synthesis within
skeletal muscle. It is phosphorylated/activated by phospho-Akt (Miyazaki and
Esser 2009). mTOR is considered to be a central sensor of multiple stimuli such
as environmental and metabolic information (i.e. amino acid availability,
extracellular milieu, ATP availability) (Kimball, Jefferson et al. 2000; Dennis,
Jaeschke et al. 2001). To investigate the role of mTOR in skeletal muscle
hypertrophy, researchers often make use of a pharmacological blocker of mTOR,
rapamyacin, to inhibit its activation. Using this approach with an in-vivo and invitro skeletal muscle model, hypertrophy was found to be blunted in both cases
(Bodine, Stitt et al. 2001; Rommel, Bodine et al. 2001). Furthermore, evidence
has shown that increases and decreases of phosphorylation at a specific mTOR
residue (Ser 2448) were associated with skeletal muscle hypertrophy and
atrophy, respectively (Reynolds, Bodine et al. 2002). In scientific models of cell
shrinkage, mTOR activation has been shown to decrease when cultured
lymphocyte cells were exposed to hyper-osmotic stress. As the cells were
returned to iso-osmotic conditions, mTOR activation returned to baseline values
(Fumarola, La Monica et al. 2005). In skeletal muscle models of cell shrinkage
(e.g. rodent hind-limb suspension), phosphorylation at Ser 2448 has been shown
to be significantly diminished (Reynolds, Bodine et al. 2002). In addition,
31 reductions of intracellular ATP levels have been shown to inhibit mTOR
activation; previous research has revealed that an extracellular hyper-osmotic
environment causes a reduction in intramuscular ATP levels (Dennis, Jaeschke
et al. 2001; Antolic, Harrison et al. 2007). Based on these observations, mTOR
phosphorylation may be decreased during hyper-osmotic stress in isolated intact
skeletal muscle.
Downstream from mTOR there are three main targets: MuRF-1/Atrogin-1,
eIF-4EBP1 and p70s6 kinase. Similar to Akt, mTOR activation has been shown
to inhibit the up-regulation of MuRF-1 and Atrogin-1, except independently of
FOXO proteins (Latres, Amini et al. 2005). Another downstream target of mTOR
is eIF-4EBP1 (eukaryotic translation initiation factor 4E-binding protein 1). eIF4EBP1 is a protein which binds to eIF-4E (eukaryotic translation initiation factor
4E) and inhibits its ability to initiate protein translation. Activated mTOR has been
shown to alleviate the inhibition of eIF-4EBP1 on eIF-4E, thereby promoting
protein translation (Hara, Yonezawa et al. 1997). The final downstream target of
mTOR is p70s6 kinase. p70s6 kinase is another serine/threonine kinase that is
involved in skeletal muscle protein synthesis. It is phosphorylated and activated
by phospho-mTOR. Following its phosphorylation, p70s6k has been shown to
phosphorylate/activate S6 ribosomal protein, which is responsible for protein
translation at the level of the ribosomes (Baar and Esser 1999). Research has
elucidated the phosphorylation of residue Thr 389 as mTOR-dependent as well
as an important phosphorylation site for optimal p70s6 kinase activity (Pearson,
Dennis et al. 1995; Burnett, Barrow et al. 1998). In models of cell shrinkage,
32 p70s6 kinase phosphorylation has been shown to be decreased. In both
lymphocyte and mouse myeloma cell culture, hyper-osmotic stress was shown to
reduce p70s6 kinase and ribosomal S6 phosphorylation (Kruppa and Clemens
1984; Fumarola, La Monica et al. 2005). In skeletal muscle models of cell
shrinkage (e.g. hind-limb suspension, microgravity) similar findings have been
observed. Spaceflight has been shown to reduce flux through the
Akt/mTOR/p70s6k pathway, while hind-limb suspension has been shown to
result in a decrease of phosphorylated p70s6 kinase (Hornberger, Hunter et al.
2001; Allen, Bandstra et al. 2009). Based on these observations, p70s6 kinase
phosphorylation levels are likely to decrease in intact isolated skeletal muscle
when placed under extracellular hyper-osmotic stress.
The Akt/mTOR/p70s6 kinase protein synthesis signalling pathway is one
of the primary pathways in skeletal muscle. Scientific models of cell shrinkage
suggest that hyper-osmotic induced cell shrinkage in skeletal muscle could lead
to decreased activity within the pathway and an overall suppression of protein
synthesis. A detailed figure of the Akt/mTOR/p70s6k protein synthesis pathway is
featured below as well as hypothesized findings of the current study (See Figure
6).
33 FIGURE 6. Summary of hypothesized findings of the Akt/mTOR/p70s6k protein
synthesis pathway, ubiquitin proteasome system, and calpain system in skeletal
muscle during extracellular hyper-osmotic stress (Adapted from (Glass 2005))
V. Summary
It has long been established that cell volume is dynamic in nature in a
variety of cell types, including skeletal muscle. Cell volume changes have been
shown to influence cellular processes, specifically metabolic activity. The theory
of cell swelling states, cellular swelling will lead to an anabolic intracellular
environment while cellular shrinkage will create a catabolic intracellular
environment. Although the purpose of these metabolic alterations is unknown,
they have been hypothesized to serve as a form of osmoregulation. Hyperosmotic induced cell shrinkage and protein turnover has been heavily researched
in hepatocytes but little work has been conducted in skeletal muscle despite its
large store of functional protein. Recently, researchers found hyper-osmotic
environment results in a suppression of total protein synthesis in muscle cell
culture (Roy 2009), however protein degradation was not measured, nor were
34 specific signalling proteins in the protein synthesis and degradation pathways. In
addition, an intact skeletal muscle model would provide a much more
physiological understanding of the effects of hyper-osmotic stress on protein
turnover.
35 CHAPTER 2: STATEMENT OF THE PROBLEM
I. Statement of the Problem
Currently, it is known that protein metabolism is altered in skeletal muscle
cell culture when exposed to extracellular hyper-osmotic stress. Specifically, an
extracellular hyper-osmotic environment results in a suppression of total protein
synthesis. However, it is unknown whether the findings in cell culture will
translate to an intact isolated skeletal muscle model. In addition, the effects of
extracellular hyper-osmotic stress on total protein degradation, ubiquitinated
protein, calpain autolysis and mTOR/p70s6k phosphorylation in skeletal muscle
are unknown.
II. Purpose
The purpose of this thesis is to:
1) Determine the influence of an extracellular hyper-osmotic environment on
overall protein synthesis and degradation in intact isolated rat skeletal muscle
2) Investigate the influence of an extracellular hyper-osmotic environment on
protein degradation pathways; specifically the ubiquitin proteasome pathway
and the calpain system.
3) Investigate the influence of an extracellular hyper-osmotic environment on the
Akt/mTOR/p70s6k protein synthesis signalling pathway.
36 III. Hypotheses
It is hypothesized that extracellular hyper-osmotic stress will:
1) Result in an overall increase in protein degradation and a suppression of
total protein synthesis.
2) Result in an up-regulation of protein degradative mechanisms, specifically:
1) in the amount of ubiquitinated proteins and 2) in the amount of µ calpain
autolysis.
3) Result in a suppression of protein synthesis signalling mechanisms,
specifically: 1) in the amount of mTOR Ser 2448 phosphorylation and 2) in
the amount of p70s6k Thr 389 phosphorylation.
37 CHAPTER 3: METHODOLOGY
I. Animals and Housing
Eight Long Evans rats (Charles River Laboratories, St. Constant, QC)
aged 4-6 weeks (127.45 ± 2.95 g), were used for total synthesis measures. An
additional eight Long Evans rats (Charles River Laboratories, St. Constant, QC)
of similar age (128.3 ± 1.94 g) were used for degradation/metabolite/intracellular
water content/Western blotting analyses (See Figure 7). Animals were housed in
groups (2-4 rats per cage) within the Brock University Animal Facility, and were
maintained on a 12:12 light-dark cycle at ~22ºC. The rats were fed a standard
rodent diet (5012 Rat diet, PMI nutrition Inc LLC, Brentwood, MO) and had ad
libitum access to food and water. All experimental procedures and protocols were
approved by the Brock University research subcommittee on animal care and
conformed to all Canadian Council on Animal Care guidelines.
FIGURE 7. Experimental Design
38 II. Osmotic Conditions Two different experimental osmotic conditions were used for muscle
incubations: a hyper-osmotic (HYPER; 400 ± 10 mmol/kg) and an iso-osmotic
(ISO; 290 ± 10 mmol/kg). These osmotic conditions have been previously
developed in our lab and have routinely been used for both soleus and extensor
digitorum longus (EDL) whole muscle incubations (Antolic, Harrison et al. 2007).
In the current study, a custom modified media based off Sigma Medium 199
(Gibco) was used for incubations. Tyrosine and phenol red were removed from
the media (see Appendix II for detailed description of Custom Media). Mannitol
was added to the media immediately prior to incubations to achieve desired
osmolalities in the HYPER group. Mannitol was used to manipulate osmolality
due to its ability to rapidly equilibrate in solution and its inability to enter
intracellular compartments (Stuart, Torres et al. 1970). The osmolality of all
incubation media was verified using a vapour pressure osmometer
(VAPRO5520, Wescor, Logan, UT).
III. Muscle Extraction The predominantly fast-twitch EDL muscle was used for incubations
because of its responsiveness to varying osmotic environments (Delp and Duan
1996; Antolic, Harrison et al. 2007). All incubations were conducted at 10-11am,
and access to chow was removed 2 hours before muscle extraction to minimize
dietary induced alterations in protein metabolism. Rats were then anesthetized
with an intraperitoneal injection of sodium pentobarbital (55 mg·kg-1) and EDL
muscles were dissected free tendon to tendon (Antolic, Harrison et al. 2007).
39 Briefly, the skin covering the ankle was cut and peeled back, exposing the leg
muscles underneath. A small incision through the fascia and superficial muscle
groups (tibialis anterior/peroneus longus) was made from the lateral side of the
patella to the lateral malleolus, exposing the EDL. The EDL was then delicately
teased out from tendon to tendon with forceps and silk sutures (4-0) were tied to
the tendons in situ. Once the muscle was properly fastened, it was excised and
quickly secured within the organ bath (< 10 sec) (Radnoti Glass Technology Inc.,
Monrovia, CA) containing approximately 15 ml of the appropriate media (See
Figure 8). The muscle was secured on one end to a hook and then hung from a
force transducer (Grass Telefactor, West Warwick, RI). The length of the muscle
was then adjusted until there was 1 g of resting tension. Organ baths were kept
at a constant temperature (30°C ± 2°C) while the media was perfused with a gas
mixture containing 95% O2, 5% CO2. 30°C ± 2°C has been previously shown to
maintain viability of similar sized muscles (Segal and Faulkner 1985; Antolic,
Harrison et al. 2007). Once both EDL muscles were removed, animals were
euthanized with an overdose of sodium pentobarbital.
40 FIGURE 8. Muscles are secured to a ‘Boomer Box’ inside the incubation bath.
The apparatus provides an attachment for the sutures tied to the muscle
(Adapted from (Antolic, Harrison et al. 2007).
IV. Protein Turn-over Measures
To determine total protein synthesis in isolated skeletal muscle during
extracellular osmotic stress, L-[14C]-U-phenylalanine uptake and incorporation
into muscle were measured. Following a 60 min pre-incubation in either ISO or
HYPER conditions, media was drained and replaced with identical media except
for the addition of 0.5mmol/L L-[14C]-U-phenylalanine (9.25MBq/L), and
subsequently incubated for an additional 75 minutes. Following the second
incubation, muscles were removed; frozen in liquid nitrogen and stored at -80°C
for future analysis.
Frozen muscles were homogenized in 0.61 mol/L trichloroacetic acid
(TCA) and centrifuged at 10,000 x g for 10 min at 4 °C to isolate the protein
fraction (Marzani, Balage et al. 2008). The resulting pellet was then solubilized in
1 mol/L NaOH at 37 °C and analyzed for 14C isotopes through scintillation
counting to determine incorporated phenylalanine (LS Multi Purpose Scintillation
41 Counter, Beckman Coulter, Mississauga, ON). Total protein synthesis was
calculated by dividing the protein-bound radioactivity by the specific activity of
free phenylalanine in the incubation medium and expressed as nmol of
phenylalanine/mg protein/75 min (Dardevet, Sornet et al. 2000).
To measure protein degradation, muscles underwent the same 60 min
pre-incubation in either HYPER or ISO media. After 60 min, the media was
replaced with an identical media; except for the addition of 0.5mmol/L
cycloheximide, muscles were then incubated for an additional 75 min.
Cycloheximide was added to the media as it prevents amino acid reincorporation into skeletal muscle tissue. Cycloheximide exerts its effect by
inhibiting the transfer of amino acids from aminoacyl tRNA to peptides at the
ribosome (Ennis and Lubin 1964). At the end of the 75 min incubation, a 2mL
aliquot of media was collected and analyzed for tyrosine concentration. The
appearance of tyrosine in the media was used as a measure of total protein
degradation, as tyrosine cannot be synthesized or degraded within muscle.
Tyrosine concentrations were determined biochemically using previously
described methods (Waalkes and Udenfriend 1957). Briefly, 1mL of 1-nitroso-2naphthol and 1mL of a nitric acid reagent were added to 2mL of sample media
where it was then stoppered, shaken and placed in a 65°C water bath for 30 min.
Upon cooling, 10mL of ethylene dichloride was added and the mixture was
shaken. The mixture was then centrifuged for 10 min at 2000rpm at 4°C and the
supernatant aqueous layer was taken and analyzed via fluorometery. The
42 sample was activated at 460nm and was measured at 570nm. To determine
tyrosine release from the muscle, samples were run against a standard curve.
V. Western Blotting Analysis:
Western blotting analyses was performed on samples to determine the
content and/or phosphorylation state of major protein synthesis and degradation
signalling pathway proteins. This was achieved through the use of antibodies
raised against the specific proteins and phospho-proteins of interest: rabbit
polyclonal mTOR (# 2972S, Cell Signalling, Danvers, MA), rabbit monoclonal
phospho-mTOR Ser2448 (#2971S, Cell Signalling, Danvers, MA), rabbit
polyclonal p70 s6 kinase α (#sc-230, Santa Cruz Biotechnology, Santa Cruz,
CA), goat monoclonal phospho-p70 s6 kinase α Thr389 (#sc-11759, Santa Cruz
Biotechnology, Santa Cruz, CA), rabbit polyclonal µ calpain (#2556S, Cell
Signalling, Danvers, MA), rabbit polyclonal ubiquitin protein conjugates
(#UG9510, Enzo Life Sciences, Farmingdale, NY) and mouse monoclonal actin
(#612656, BD Biosciences, Mississauga, ON). Ubiquitin conjugates were
analyzed in relation to total actin where as phospho-p70 s6 kinase α Thr389 and
phospho-mTOR Ser2448 were analyzed in relation to total p70s6 kinase and
total mTOR, respectively. Autolysis of µ calpain was represented as the
increasing appearance of the lower bands at 78 and 76 kDa while the higher 80kDa band represents un-autolyzed µ calpain. Each antibody was tested prior to
analysis to optimize protein load.
Samples were homogenized by hand in an ice cold buffer (250mM
sucrose, 100mM KCl, 10mM EGTA) with the addition of a protease inhibitor
43 (#11836170001, Roche, Laval, QC) and phosphatase inhibitor (#04906837001,
Roche, Laval, QC). Following homogenization, a Bradford assay was run to
determine protein concentration and samples were then diluted with H2O and a
sample buffer ( 0.5M Tris-HCl, 2mL glycerol, 10% SDS, 1% bromophenol blue,
1mL β-mercaptoethanol, and dH2O) to a final protein concentration of 1 GAWul-1.
Standard SDS-PAGE electrophoresis was performed with the use of 7.5%
(mTOR, phospho-mTOR, µ calpain and ubiquitin protein conjugates) or 10% (p70
s6 kinase, phospho-p70 s6 kinase) separating gel and loaded with 8 - 20 µl of
protein. Molecular weight standards were also included alongside samples on
each gel (#161-0375, BioRad, Mississauga, ON). Samples were run for
anywhere from 75-120 minutes at 100-160V depending on the specific protein
(See Appendix). Electrophoretically separated proteins were then transferred to
either nitrocellulose membrane (µ calpain) or polyvinylidene fluoride membrane
(PVDF) (mTOR, phospho-mTOR, ubiquitin protein conjugates, p70 s6 kinase and
phospho-p70 s6 kinase) for 75-120 minutes at 100V in a transfer buffer (1.25%
SDS, 312.5mM Tris Base, 2.4M glycine, 20% methanol). Following transfer,
membranes were blocked in a blocking solution to prevent non-specific binding,
in either 5% (w/v) BSA (bovine serum albumin) or 5% (w/v) non-fat dry milkTBST solution (TBST buffer: 20mM tris base, 137mM NaCl, and 0.1% (v/v)
tween 20, pH 7.5) for 60 minutes at room temperature. Membranes were then
washed 5 times for 5 minutes each with TBST buffer. Following this series of
washes, membranes were incubated overnight with the appropriate antibody
dilution. Dilutions were determined via pilot work. Antibodies were placed in
44 either a 1-5% milk TBST solution or a 1% (w/v) BSA in PBST solution (PBST
buffer: 137mM NaCl, 3mM KCl, 10mM Na2HPO4, 2mM KH2PO4, 0.05% (v/v)
tween 20, pH 7.2) depending on protein. Membranes were rocked overnight at
4°C. Post primary antibody incubation, membranes were again washed 5 times
for 5 minutes each with TBST buffer to remove unbound antibody. Following
these washes, membranes were incubated for 1 hour at room temperature with
an appropriate HRP conjugated secondary antibody at the appropriate dilution.
This was followed by another series of washes in TBST buffer (5 X 5min).
Membranes were then treated with Chemi Glow West Substrate (#WBKLS0500,
Billerica, MA) for 5 minutes, developed using the FluroChem imaging system
(Alpha-Innotec, Santa Clara, CA), and quantified using the AlphaEaseFC
Software (Alpha-Innotec, Santa Clara, CA). Band density was determined with
“Image J” software (National Institutes of Health, Bethesda, MD).
VI. Metabolite Analysis:
For metabolite analysis, a small portion of frozen muscle was chipped off,
dehydrated, powdered and removed of any connective tissue or blood. The
powdered sample was then acid extracted for quantification of; adenosine
triphosphate (ATP) and phospho-creatine (PCr). Previously described
flourometric techniques were used to analyze the concentrations of the
metabolites (Harris, Hultman et al. 1974; Green, Thomson et al. 1987). To assure
accuracy and reliability of the measures, each sample was analyzed in triplicate
during the same experimental session.
45 VII. Statistical Analysis
All data were analyzed using independent samples t tests to compare the
means of the two experimental groups; ISO and HYPER. The level of statistical
significance was set at p≤0.05 for all analysis. All statistical analyses were
conducted using GraphPad Prism 5 (La Jolla, CA) and all values are expressed
as the means ± standard error (SE).
46 CHAPTER 4: RESULTS
I. General Observations
No statistical differences were observed between the weights of the rats in
each experimental condition (see Table 1). Media osmolality measured prior to
incubation fell within ± 10 GGIFWEA-1 of the respective target osmolalities. At this
time, the HYPER condition osmolality was found to be significantly (p<0.05) higher than the ISO condition.
Table 1: Rat Weights and Media Osmolalities
Experimental Group
Rat Weight (g)
Osmolality
ISO
125.1 ± 1.7
285.4 ± 1.2
HYPER
130.6 ± 2.9
394.3 ± 0.7*
Note: Values are expressed as mean ± SE (n = 16 muscles per group). *
Significantly different (p<0.05) from ISO.
II. Relative Water Content
Following the 60 min pre-incubation and 75 min incubation, relative water
content of the muscle was found to be significantly (p<0.05) decreased (4.1%) in
the HYPER condition as compared to ISO (see Figure 9).
47 Water Content (%)
80
78
76
74
72
70
*
ISO
HYPER
FIGURE 9. EDL water content during hyper-osmotic stress. Values are
expressed as means ± SE (n = 7 per group). * Significantly different (p<0.05)
from ISO.
III. Metabolites
In the EDL muscle, both ATP and PCr concentrations were significantly
(p<0.05) lower in the HYPER condition compared to ISO (see Table 2).
Table 2: High Energy Phosphate Metabolite Concentrations
Experimental Group
ATP
PCr
ISO
32.1 ± 0.4
77.7 ± 0.4
HYPER
30.0 ± 0.6*
37.2 ± 0.6*
Note: All values were derived from EDL muscles. All values expressed in
GGIFWEA-1 dry wt. n = 7 for each group. ATP = adenosine triphosphate, PCr =
phosphocreatine. * Significantly different (p<0.05) from ISO.
IV. Protein Synthesis
Incorporated phenylalanine was found to be significantly (p<0.05) lower
(41%) in the HYPER condition compared to the ISO control (See Figure 10).
48 Incorporated Phenylalanine
(nmol/mg/75 min)
15
10
*
5
0
ISO
HYPER
FIGURE 10. Incorporated phenylalanine in skeletal muscle during iso-osmotic
and hyper-osmotic stress. Values are expressed as means ± SE (n = 8 per
group). * Significantly different (p<0.05) from ISO.
V. Protein Degradation
Tyrosine release was found to be significantly (p<0.05) increased (43%)
in the HYPER condition compared to ISO (see Figure 11).
Tyrosine Release
(nmol/mL)
2.5
*
2.0
1.5
1.0
0.5
0.0
ISO
HYPER
FIGURE 11. Protein degradation as estimated by tyrosine release from skeletal
muscle during iso-osmotic and hyper-osmotic stress. Values are expressed as
means ± SE (n = 8 per group). * Significantly different (p<0.05) from ISO.
49 VI. Western Blotting Analyses
Total mTOR protein content was found to be similar in both experimental
groups (see Figure 12). However, the amount of mTOR phosphorylation in
relation to total mTOR was found to be significantly (p<0.05) increased (15%) in
the HYPER condition (see Figure 13). Total p70s6 kinase protein content was
found to be similar in both experimental groups (see Figure 14) while p70s6
kinase phosphorylation in relation to total p70s6 kinase content was found to be
significantly (p<0.05) decreased (23%) in the HYPER condition compared to ISO
control (see Figure 15).
Total actin in skeletal muscle was found to be consistent between the ISO
and HYPER groups (see Figure 16). The amount of ubiquitinated protein in
relation to total actin was also found to be consistent between the ISO and
HYPER conditions (see Figure 17). Unautolyzed µ calpain (80kDa) was found to
be significantly (p<0.05) decreased in the HYPER condition compared to ISO
control (see Figure 18). Autolyzed µ calpain (78/76 kDa) was found to be
significantly (p<0.05) increased in HYPER compared to the ISO control (see
Figure 18).
50 2.5
Total mTOR
2.0
1.5
1.0
0.5
0.0
ISO
HYPER
FIGURE 12. Total mTOR protein content in skeletal muscle during iso-osmotic
and Hyper-osmotic Stress. Values are expressed as means ± SE (n = 8 per
group).
Phosphorylated mTOR/Total mTOR
1.5
*
1.0
0.5
0.0
ISO
HYPER
FIGURE 13. mTOR phosphorylation in relation to total mTOR in skeletal muscle
during iso-osmotic and hyper-osmotic Stress. Values are expressed as means ±
SE (n = 8 per group). * Significantly different (p<0.05) from ISO.
51 Total p70s6 kinase
1.5
1.0
0.5
0.0
ISO
HYPER
FIGURE 14. Total p70s6 kinase content in skeletal muscle during Iso-osmotic
and Hyper-osmotic Stress. Values are expressed as means ± SE (n = 8 per
group).
Phosphorylated p70s6k/Total p70s6k
1.0
0.8
*
0.6
0.4
0.2
0.0
ISO
HYPER
FIGURE 15. p70s6 kinase phosphorylation in relation to total p70s6 kinase in
skeletal muscle during iso-osmotic and hyper-osmotic stress. Values are
expressed as means ± SE (n = 8 per group). * Significantly different (p<0.05)
from ISO.
52 Total Actin
1.5
1.0
0.5
0.0
ISO
HYPER
FIGURE 16. Total actin protein content in skeletal muscle during iso-osmotic and
hyper-osmotic stress. Values are expressed as means ± SE (n = 4 per group).
Ubiquitinated Protein/Total Actin
0.4
0.3
0.2
0.1
0.0
ISO
HYPER
FIGURE 17. Total ubiquitinated protein in relation to total actin in skeletal muscle
during iso-osmotic and hyper-osmotic stress. Values are expressed as means ±
SE (n = 4 per group).
53 ←78/76kDa 2.0
80kDa
78/76kDa
1.5
α
1.0
*
0.5
R
0.0
H
YP
E
IS
O
Relative Density of Bands
80kDa→ FIGURE 18. Unautolyzed (80kDa) and autolyzed (78/76kDa) calpain in skeletal
muscle during iso-osmotic and hyper-osmotic stress. Values are expressed as
means ± SE (n = 6 per group). α Significantly different (p<0.05) from ISO –
80kDa. * Significantly different (p<0.05) from ISO – 78/76kDa.
54 CHAPTER 5: DISCUSSION
Many studies have examined the effect of hyper-osmotic stress on protein
turnover in whole body and hepatocyte models, but few have examined the effect
in skeletal muscle (Haussinger, Hallbrucker et al. 1991; Stoll, Gerok et al. 1992;
Berneis, Ninnis et al. 1999; Roy 2009). Furthermore, the few studies that have
investigated skeletal muscle used varying models. To the authors knowledge the
current study is the first to have utilized used an isolated intact skeletal muscle
model. Given that this is the first known study to have used this particular model
to examine protein turnover under extracellular hyper-osmotic stress,
comparative literature is limited.
The primary finding of the current study was the influence of extracellular
hyper-osmotic stress on protein turnover in skeletal muscle. Extracellular hyperosmotic stress was shown to suppress protein synthesis as well as stimulate
protein degradation. The expression of major protein synthesis and degradation
signalling pathway proteins were generally in agreement with the protein turnover
data. Contrary to our hypotheses, mTOR Ser 2448 phosphorylation was found to
increase while ubiquitin conjugates were unchanged during extracellular hyperosmotic stress. In support of our hypotheses, p70s6 kinase Thr 389
phosphorylation was decreased while µ calpain autolysis was increased during
extracellular hyper-osmotic stress. Overall, these results suggest that osmotically
induced decreases of cell volume influence protein turnover in an intact skeletal
muscle model and promote an intracellular protein catabolic environment.
55 I. General Observations
Intact isolated skeletal muscle that is exposed to extracellular hyperosmotic stress has been shown to result in a decline of cell volume (Antolic,
Harrison et al. 2007; Mulligan 2008). These findings were confirmed in the
current study. The 60 minute pre-incubation and 75 minute incubation in hyperosmotic media resulted in a 4.1% decrease in intracellular water content (p<0.05)
compared to the iso-osmotic control. The intracellular water loss observed in the
current study was similar to previous studies within our lab despite an additional
75 minutes of incubation time (Antolic, Harrison et al. 2007; Mulligan 2008). This
suggests that RVI processes were likely complete by 60 minutes of incubation
time and the incubated muscles were at a new stable equilibrium volume.
The present work also demonstrated a significant (p<0.05) decrease of
intramuscular ATP and PCr concentrations during extracellular hyper-osmotic
stress. These findings were in agreement with previous work conducted within
our lab (Antolic, Harrison et al. 2007). The reduction of intramuscular ATP and
PCr concentrations could be attributed to several factors. First of all, the
macromolecular crowding experienced during hyper-osmotic induced cell
shrinkage, known to influence intracellular protein activity, could negatively
influence energy producing pathways within the cell (Garner and Burg 1994).
Another possible explanation involves the theory of cell swelling. The
degradation of intracellular molecules increases intracellular osmolality, therefore
the breakdown of ATP and PCr may have been used as an acute volume
regulatory mechanism to help increase intracellular osmolality and prevent
56 further water loss (Antolic, Harrison et al. 2007). Finally, the reduction of
intramuscular ATP and PCr concentrations could have been the result of
increased energy demand. There are three main consumers of ATP in skeletal
muscle: Na+/K+ ATPase, Ca2+ ATPase and acto-myosin ATPase. Since the
muscle was not producing force, it is likely that acto-myosin ATPase was not a
significant user of ATP during the experiment. The primary role of Na+/K+
ATPase is to maintain an electrochemical gradient at the sarcolemma but it also
serves as an indirect form of osmoregulation. During RVI, NKCC1 is activated to
transport 2 Na+, K+ and Cl- into the cell in an attempt to increase intracellular
osmolality; it is hypothesized that the Na+/K+ transporter is activated shortly after
to maintain the electrochemical gradient in spite of the mass ion entry (Lindinger,
Hawke et al. 2002). Ca2+ ATPase is also a candidate for increased activity during
hyper-osmotic stress due to the phenomenon of Ca2+ sparks. Research has
shown that extracellular hyper-osmotic stress results in transient increases of
intracellular Ca2+ concentration in skeletal muscle (Apostol, Ursu et al. 2009). It is
suggested that these Ca2+ sparks are the result of a mechanical disruption
between DHPR and ryanodine receptors during cell shrinkage. With this
suspected increase of intramuscular Ca2+ concentration during extracellular
hyper-osmotic stress, Ca2+ ATPase activity could be increased as it works to
transfer Ca2+ from the cytosol to the SR. In summary, ATP and PCr
concentrations were likely reduced due to an increased energy demand within
the muscle during extracellular hyper-osmotic stress. It is likely due to multiple
57 sources, specifically increased activity of Na+/K+ ATPase and Ca2+ ATPase due
to volume regulatory mechanisms and Ca2+ sparks, respectively.
II. Protein Synthesis
In the current study, extracellular hyper-osmotic stress was shown to
result in a significant (p<0.05) decrease of phenylalanine incorporation in skeletal
muscle. The suppression of protein synthesis, as estimated by phenylalanine
incorporation, was in accordance with findings in hepatocyte and skeletal muscle
cell culture models (Stoll, Gerok et al. 1992; Roy 2009). These results lend
support to the theory of cell swelling as they are indicative of an intracellular
protein catabolic environment. In this case, hyper-osmotic induced cell shrinkage
appears to trigger an intracellular signal cascade which eventually leads to an
inhibition of protein synthesis in skeletal muscle. The intracellular signalling
mechanism responsible for this suppression of protein synthesis was largely
unknown but advancements have been made. Research has elucidated p70s6
kinase activity as an important contributor to the suppression of protein synthesis
during hyper-osmotic stress in both myeloma and lymphocyte cell culture
(Kruppa and Clemens 1984; Fumarola, La Monica et al. 2005). In addition,
investigators have established an association between p70s6 kinase
phosphorylation at Thr 389 and hypertrophy and atrophy in skeletal muscle (Baar
and Esser 1999; Terzis, Georgiadis et al. 2008). However, the effects of
extracellular hyper-osmotic stress on p70s6 kinase activity in skeletal muscle
were previously unknown. In the current study, it was determined that hyperosmotic induced cell shrinkage significantly (p<0.05) decreased p70s6 kinase
58 phosphorylation at Thr 389. In addition to the aforementioned literature, these
findings are also in agreement with skeletal muscle models of cell shrinkage;
rodent hind-limb suspension and micro-gravity models have both been shown to
result in a decrease of p70s6 kinase phosphorylation (Hornberger, Hunter et al.
2001; Allen, Bandstra et al. 2009). Based on these observations, it is plausible
that the decrease of p70s6 kinase phosphorylation at Thr 389 could be
contributing to the suppression of protein synthesis observed in skeletal muscle
during hyper-osmotic induced in muscle cell shrinkage.
The observed increase of mTOR phosphorylation at Ser 2448 during
extracellular hyper-osmotic stress in skeletal muscle was not an expected
finding. It was originally believed that mTOR phosphorylation/activation would be
reduced due to its known role in skeletal muscle hypertrophy as well as the
observed decrease in protein synthesis and p70s6 kinase phosphorylation within
the current study (Bodine, Stitt et al. 2001). However, the role of mTOR in
skeletal muscle is very diverse; it is considered to be a central sensor with many
intracellular influences. There are five main upstream effectors of mTOR activity
in skeletal muscle: growth factors (e.g. insulin, IGF-1), mechanical stress, energy
status (e.g. ATP concentrations, AMPK), cellular stress (e.g. hypoxia) and amino
acid availability (Bolster, Jefferson et al. 2004; Foster and Fingar 2010). Growth
factors, mechanical overload and sufficient amino acid availability are considered
to be activators of mTOR in skeletal muscle, while reductions of intracellular ATP
and cellular stress inhibit its action (Bolster, Jefferson et al. 2004; Foster and
Fingar 2010). In the current study, growth factors (e.g. insulin and IGF-1) were
59 absent from the experimental media and isolated EDL muscles were not
stimulated, therefore it seems unlikely that either could be cause for increased
mTOR phosphorylation in skeletal muscle during hyper-osmotic stress. However,
the apparent intracellular protein catabolism suggests that the skeletal muscle
amino acid pool may be enlarged during extracellular hyper-osmotic stress.
Although the molecular pathway in which mTOR senses and responds to amino
acid availability is poorly understood, research has consistently shown that an
increase of amino acid availability results in an increase of mTOR
phosphorylation at Ser 2448 in skeletal muscle (Anthony, Anthony et al. 2001;
Reynolds, Bodine et al. 2002; Bolster, Vary et al. 2004). Furthermore, some
researchers have hypothesized that amino acid availability is the dominant input
of mTOR (Hara, Yonezawa et al. 1998; Beugnet, Tee et al. 2003; Nobukuni,
Joaquin et al. 2005).
During hyper-osmotic induced cell shrinkage, the skeletal muscle amino
acid pool has been shown to be used as a means of osmoregulation (Lang,
Busch et al. 1998; Hyde, Taylor et al. 2003). During RVI, the acute accumulation
of ions can disrupt intracellular processes (e.g. sarcolemmal gradient, cytosolic
pH) therefore amino acids are accumulated in the long-term to maintain cell
function and promote water retention. However, not all amino acids exert the
same influence on cell volume. Glutamine, glutamate, glycine, aspartate and
taurine have been shown to have the greatest influence on cell volume (Lang,
Busch et al. 1998). Amino acids are moved across the cell membrane by way of
4 known skeletal muscle amino acid transporter systems: A, ASC, L and Nm
60 (Bonadonna, Saccomani et al. 1993). Each system is responsible for the
movement of several amino acids across the membrane. In addition, all
transporters, with the exception of L system, are Na+-dependent and carry Na+
across the membrane with amino acids. In terms of osmoregulation, research
has shown that hyper-osmotic stress results in an increased expression of the
System A amino acid transporter (Hyde, Taylor et al. 2003). The system A amino
acid transporter is primarily responsible for the movement of glutamine, alanine,
threonine, asparagine, serine and methionine (Bonadonna, Saccomani et al.
1993; Hyde, Taylor et al. 2003). Based on these observations, an increase of the
intracellular amino acid pool could be cause for the increase of mTOR
phosphorylation at Ser 2448 in skeletal muscle during extracellular hyper-osmotic
stress.
Further upstream the signalling mechanism responsible for the
suppression of protein synthesis during extracellular hyper-osmotic stress is
much less clear. Researchers hypothesize that macromolecular crowding may
play a role in detection of cell volume as well as in the initiation of metabolic
signalling response (Haussinger 1996). It is believed that cellular shrinkage alters
the concentration of cytosolic proteins (e.g. protein kinases and phosphatases)
and subsequently determines their activity by influencing the tendency to
associate with their substrates. This is believed to eventually result in volumeregulatory responses as well as changes to cellular metabolism and gene
expression. Research in hepatocytes has focused on MAP kinase regulation
during hyper-osmotic stress as a potential link between cell volume detection and
61 protein synthesis but additional research is necessary, particularly in skeletal
muscle (Haussinger 1996). The activation of protein degradation pathways may
also play a role in the regulation of protein synthesis during hyper-osmotic stress
as the liberation of amino acids could potentially influence protein synthesis
activity.
III. Protein Degradation
In the current study, extracellular hyper-osmotic stress was shown to
result in a significant (p<0.05) increase of protein degradation in skeletal muscle,
as estimated by tyrosine release. These findings are in agreement with previous
research in hepatocytes (Haussinger, Hallbrucker et al. 1991). Similar to protein
synthesis findings, the increase of increased protein degradation is consistent
with the theory of cell swelling. In this case, hyper-osmotic induced cell shrinkage
creates an intracellular protein catabolic environment and leads to the
degradation of intracellular proteins into amino acids. However, there has also
been a lack of literature investigating the effect of extracellular hyper-osmotic
stress on the activation of protein degradation mechanisms in skeletal muscle.
There are two major mechanisms of protein degradation within skeletal muscle:
the ubiquitin proteasome system and calpain system (Jackman and Kandarian
2004). In this study, ubiquitin conjugates were found to be consistent between
both experimental groups while µ calpain was found to have a significantly
(p<0.05) higher percentage of autolysis in the HYPER condition.
Based on skeletal muscle models of cell shrinkage (e.g. hind-limb
suspension and space flight), extracellular hyper-osmotic stress was expected to
62 result in an increase in ubiquitinated protein (Bodine, Latres et al. 2001; Ikemoto,
Nikawa et al. 2001). One reason that we may have not seen an increase in
ubiquitin conjugates is due to the time course of action of the ubiquitin
proteasome system. The ubiquitin proteasome system is thought to be a longer
term mechanism of protein degradation, therefore our incubation may not have
been long enough in duration to produce a change in ubiquitin conjugates (Attaix,
Aurousseau et al. 1998). Another potential explanation for the ubiquitin
conjugates results involves a potential relationship between protein synthesis
signalling pathways and the ubiquitin proteasome system. Research has
suggested that cross-talk can occur between protein synthesis signalling
pathways and E3 ligases in skeletal muscle. E3 ligases that are responsible for
poly-ubiquitination and it is not until at least four ubiquitin molecules are attached
to a protein that it can be degraded by the proteasome (Pickart 2000). To
investigate a potential relationship between the Akt/mTOR/p70s6 kinase protein
synthesis pathway and E3 ligases, researchers used dexamethasone and IGF-1
to induce skeletal muscle atrophy/cell shrinkage and hypertrophy/cell swelling,
respectively (Stitt, Drujan et al. 2004). During dexamethasone treatment, FOXO
proteins were activated and lead to the up-regulation of E3 ligases. Conversely,
during IGF-1 treatment, FOXO proteins and E3 ligases were inhibited (Sandri,
Sandri et al. 2004). In addition, it was shown that increased activity of the
PI3K/Akt/mTOR pathway could block the atrophy-inducing effects of
dexamethasone, specifically the up-regulation of E3 ligases (Stitt, Drujan et al.
2004). These observations demonstrated that activity of protein synthesis
63 pathways can influence the activity of protein degradation pathways, specifically
E3 ligases. To further investigate this relationship, researchers used rapamyacin
to inhibit the action of mTOR in IGF-1 treated muscles. This action eliminated
some of the IGF-1-mediated transcriptional changes, specifically the downregulation of E3 ligases (Latres, Amini et al. 2005). This action occurred
independently of FOXO proteins. These observations suggest that mTOR plays a
role in the down-regulation E3 ligases. In the current study, extracellular hyperosmotic stress resulted in an increase in mTOR phosphorylation. Therefore, it is
possible that the increase of mTOR activation inhibited an up-regulation of
ubiquitin conjugates. Future research should investigate changes in mRNA
expression of E3 ligases, Atrogin-1 and MuRF-1, in a similar experimental model
to investigate the acute activation of the ubiquitin proteasome system.
The increase in calpain autolysis was an expected finding due to the
known presence of Ca2+ sparks during hyper-osmotic stress in skeletal muscle
(Apostol, Ursu et al. 2009). Calpain autolysis is closely associated with activation
of calpain therefore an increase in autolysis is representative of its degradative
activity (Baki, Tompa et al. 1996). Previous literature in skeletal muscle has
shown that extracellular hyper-osmotic stress results in spontaneous transient
increases in intracellular Ca2+ concentration (Apostol, Ursu et al. 2009). Based
on the Ca2+-dependent regulation of calpain, these observations suggest that
Ca2+ sparks are responsible for the increase of calpain autolysis. The
concentration of these sparks is currently unknown but are assumed to reach the
activation threshold of µ calpain, 3-50 µm (Goll, Thompson et al. 2003). It has
64 been suggested that hyper-osmotic stress disrupts the structural interaction
between DHPR and ryanodine receptors to allow Ca2+ release from the SR.
However, further research should be conducted to verify this hypothesis.
Despite the findings of this study, there is a lack of understanding
regarding the signalling mechanisms which link hyper-osmotic induced cell
shrinkage to protein degradation mechanisms in skeletal muscle. Previous work
in hepatocytes has hypothesized the microtubule network may be a key
component in the regulation of proteolysis. In contrast to declines in cell volume,
cellular swelling has been shown to inhibit proteolysis in hepatocytes in the
theory of cell swelling. Researchers have attempted to elucidate the mechanisms
underlying these findings through the induction of cell swelling with hypo-osmotic
media, insulin and glutamine. Each of these agents were shown to increase
tubulin mRNA levels 1.9, 2.1 and 2.7 fold, respectively (Haussinger, Stoll et al.
1994). These findings lead researchers to believe that cellular swelling creates a
stabilizing environment on microtubules. In a follow-up study, perfused
hepatocytes were again exposed to hypo-osmotic media, insulin and glutamine
but also colchicine (vom Dahl, Stoll et al. 1995). Colchicine is a microtubule
inhibitor which binds to tubulin and prevents polymerization without affecting cell
volume (Ai, Ralston et al. 2003). In this study, colchicine was found to abolish the
inhibition of hepatic proteolysis induced by cell swelling in each case.
Furthermore, perfused hepatocytes were supplied with anti-proteolytic amino
acids which do not influence cell volume (e.g. asparagine), as well as colchicine,
and proteolysis inhibition was found to be unaffected (vom Dahl, Stoll et al.
65 1995). This suggests that the regulation of proteolysis by way of cell volume
alterations requires an intact microtubule network. During conditions of cell
shrinkage, it seems likely that the microtubular network could become disrupted,
potentially resulting in the activation of an intracellular signalling cascade
promoting proteolysis. Although this series of studies has begun to elucidate the
link between cell volume and proteolysis, the picture is still unclear. Future work
should focus on establishing the mechanism responsible for control of these
processes in other tissues such as skeletal muscle.
Protein degradation, as estimated by intramuscular tyrosine release, was
significantly (p<0.05) increased during extracellular hyper-osmotic stress in
skeletal muscle. This increase may be related to the activation of the calpain
system. These findings are in agreement with the theory of cell swelling but
cannot be proven until research elucidates the mechanisms which link cell
shrinkage to protein degradation pathways. Future research should also focus on
confirming the regulation of E3 ligases by protein synthesis signalling proteins.
IV. Summary and Conclusions
Cell volume regulation is an extremely complex process with many
underlying mechanisms. The link between cell volume and metabolic activity has
been studied extensively and led to the theory of cell swelling. The theory of cell
swelling states that cell volume acts as intracellular signal which influences
metabolic pathways in the cell. Cell swelling results in an increase in anabolic
activity, while cellular shrinkage leads to a catabolic cellular environment.
Whether these metabolic fluctuations are directly related to cell volume regulation
66 or an unintended consequence remains unclear. In support of osmoregulation,
the osmotic effect of individual amino acids is far greater than that of synthesized
proteins; therefore inhibiting protein synthesis could theoretically increase
intracellular osmolality, which in turn would promote water retention. However, it
is difficult to classify these metabolic fluctuations as a means of osmoregulation
for several reasons. First of all, not every effect of cell volume change on
intracellular signalling is related to cell volume regulation. Secondly, different cell
types will make use of different osmoregulation systems; therefore an
established mechanism in one tissue must be confirmed in all other types.
Another difficulty arises due to the multiple mechanisms that cells likely use in
parallel to regulate cell volume. These systems will often overlap which makes it
very difficult to analyze the individual effect of intracellular signalling mechanisms
on osmoregulation (Lang, Busch et al. 1998). Overall, an intracellular mechanism
serves osmoregulation if it is modified by alterations in cell volume and a
qualitative/quantitative modification of this intracellular mechanism triggers the
appropriate alterations of cell volume. For all these reasons, it is difficult to
characterize the suppression of protein synthesis as a form of osmoregulation
despite it being an attractive hypothesis.
The results of this study are extremely novel as this is the first known
study to examine hyper-osmotic induced cell shrinkage and protein turnover in
intact skeletal muscle. The most important finding in this study was that exposure
to hyper-osmotic stress lead to the suppression of protein synthesis and the
stimulation of protein degradation in intact isolated skeletal muscle. This
67 suggests that cellular shrinkage creates an intracellular catabolic environment,
but as previously mentioned, the signaling mechanisms which link cell volume
detection to protein turnover must be established further to verify this. The
second part of this study aimed to establish some of the signaling mechanisms
involved. Within protein synthesis, mTOR phosphorylation was found to be
significantly (p<0.05) increased while p70s6 kinase phosphorylation was found to
be significantly (p<0.05) decreased. This suggests that p70s6 kinase may be
involved in the suppression of protein synthesis while mTOR may be primarily
regulated by nutrient availability. Within protein degradation, calpain autolysis
was found to be significantly (p<0.05) increased while the ubiquitin proteasome
system experienced no change. This suggests that calpain activity likely
contributes to the increase in protein degradation during hyper-osmotic stress
while the ubiquitin proteasome system may be influenced by mTOR activation. A
summary of the protein turnover signaling findings in the current study can be
seen below (see Figure 19).
68 FIGURE 19. Summary of protein turnover data in skeletal muscle during isoosmotic and hyper-osmotic stress.
V. Future Directions
It is clear that skeletal muscle protein metabolism is altered in response to
hyper-osmotic stress. However, it is unclear as to whether these perturbations
are part of a volume regulatory response, or are simply a consequence of the
changes in cell volume. The current study has begun to establish the signaling
proteins involved in protein synthesis and degradation pathways during hyperosmotic stress, but the bigger picture remains incomplete. In order to help
determine this, future studies should attempt to elucidate the remainder of the
signaling proteins involved, particularly upstream. In addition, future research
should re-investigate the activation of the ubiquitin proteasome system during
hyper-osmotic stress in skeletal muscle. Using an upstream measurement such
as mRNA expression of muscle specific E3 ligases (i.e. Atrogin-1 and MuRF-1)
would allow for an acute determination of its activation during hyper-osmotic
induced cell shrinkage in skeletal muscle. Lastly, future research should attempt
69 to replicate these findings within an in-vivo skeletal muscle model. Through the
use of intravenous tracer infusion and muscle biopsy techniques, muscle cell
volume changes could be achieved and protein metabolism quantified.
Conducting this research could potentially facilitate new opportunities to
investigate clinical conditions where decreases in muscle cell volume are an
issue.
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85 APPENDIX I: BREAKDOWN OF MODIFIED SIGMA MEDIA-199
Components
M4530
[1X]
AW)-1
Inorganic Salts
CaCl2·2H2O
Fe(NO3)3·9H2O
MgSO4 (anhyd)
KCl
KH2PO4
Na·Acetate (anhyd)
NaHCO3
NaCl
Na2HPO4 (anhyd)
NaH2PO4 (anhyd)
0.265
0.00072
0.9767
0.4
-0.05
2.2
6.8
-0.122
Amino Acids
DL-Alanine
L-Arginine·HCl
DL-Aspartic Acid
L-Cysteine·HCl·H20
L-Cystine·2HCl
DL-Glutamic Acid
L-Glutamine
Glycine
L-Histidine·HCl·H2O
Hydroxy-L-Proline
DL-Isoleucine
DL-Leucine
L-Lysine·HCl
DL-Methionine
DL-Phenylalanine
L-Proline
DL-Serine
DL-Threonine
DL-Tryptophan
DL-Valine
0.05
0.07
0.06
0.00011
0.026
0.1336
0.1
0.05
0.02188
0.01
0.04
0.12
0.07
0.03
0.05
0.04
0.05
0.06
0.02
0.05
86 Vitamins
Ascorbic Acid·Na
D-Biotin
Calciferol
Choline Chloride
Folic Acid
Menadione (sodium bisulfite)
myo-Inositol
Niacinamide
Nicotinic Acid
p-Amino Benzoic Acid
D-Pantothenic Acid·½Ca
Pyridoxal·HCl
Pydridoxine·HCl
Retinol Acetate
Riboflavin
DL-α-Tocopherol Phosphate·Na
Thiamine·HCl
0.000056
0.00001
0.0001
0.0005
0.00001
0.000016
0.00005
0.000025
0.000025
0.00005
0.00001
0.000025
0.000025
0.00014
0.00001
0.00001
0.00001
Other
Adenine Sulfate
Adenosine Triphosphate·2Na
Adenosine Monophosphate·Na
Cholesterol
Deoxyribose
Glucose
Glutathione (reduced)
Guanine·HCl
HEPES
Hypoxanthine
Tween 80
Ribose
Thymine
Uracil
Xanthine·Na
0.01
0.001
0.000238
0.0002
0.0005
1.0
0.00005
0.0003
-0.0003
0.02
0.0005
0.0003
0.0003
0.000344
87 APPENDIX II: LABORATORY PROCEDURES
I. Metabolite Protocol
I. Metabolite Extraction
Make up 0.5M PCA solution (21.5mL 70% PCA into 500mL dH2O).
Make up 2.3M KHCO3 (2.3g KHCO3 into 10mL dH2O). Make fresh daily.
Freeze dry tissue (overnight to ensure all water is removed)
Store with dry rite in freezer until powdering
Tease out connective tissue and powder
Place in pre-weighed microcentrifuge tube and weigh (3-5mg)
Place tubes in an ice bucket (make sure tubes remain cold)
Add 600 µL of pre-cooled 0.5 M PCA
Extract for 10 minutes, vortexing several times (ensure all tissue is in contact
with PCA)
10) Centrifuge for 10 minutes at 15 000 G (spinning helps remove some of the
enzymes that can influence concentration)
11) Remove 540 µL and place in freezer (-20ºC) for 10 min
12) To the frozen supernatant add 135 µL of 2.3 M KHCO3 and vortex until liquid
(addition of KHCO3 to a frozen supernatant prevents foaming over)
13) Centrifuge 10 min 0ºC at 15 000G. Remove supernatant to assay
metabolites.
1)
2)
3)
4)
5)
6)
7)
8)
9)
II. Muscle Adenosine Triphosphate (ATP) and Phosphocreatine (PCr) Assay
CREATINE KINASE
P-CREATINE + ADP--------------------------------------- CREATINE + ATP
HEXOKINASE
ATP + GLUCOSE---------------------------------------- ADP + GLUCOSE-6-P
G-6-P-DH
GLUCOSE-6-P + NADP------------------------------- GLUCANOLACTONE +
NADPH
88 Reagent
STOCK
CONC.
FINAL
CONC.
VOLUME
25ML
VOLUME
50ML
VOLUME
100ML
1.00 M
50 mM
1.25 ml
2.5 ml
5.00 ml
2. MgCl2 (on
shelf) fresh
(.2033g/ml)
1.00 M
1.0 mM
25.00 µL
50.00 µL
100.00 µL
3. D.T.T.
0.5 M
0.5 mM
25.00 µL
50.00 µL
100.00 µL
100.0
mM
100.0
µM
25.00 µL
50.00 µL
100.00 µL
50.0 mM
50.0 µM
25.00 µL
50.00 µL
100.00 µL
2660
U/ml
0.02
U/ml
1.00 µL
2.00 µL
4.00 µL
1. Tris (on shelf)
(pH 8.1)
stored in fridge
(found in -20)
aliquots -80
4. Glucose
(on shelf)
aliquots
( -80)
5. NADP
(found in -20)
aliquots
( -80)
6. G-6-P-DH
(found in fridge)
Sigma (G-5760)
7. ADP (found
Solid
in -20)
Sigma (A-2754)
8. Creatine
Kinase
(found in -20)
324 U/mg
Sigma (C-3755)
89 Note: Mix reagents 1-5 together. Bring to volume with distilled water and adjust
to pH 8.1. Then add reagent 6. Mix by inversion with enzymes.
Preparation of Dilute Enzyme
For step 2. Mix 2.5 µL of Hexokinase with 1ml of buffer. Mix by inversion.
For step 3. Mix ~1.5 mg of phosphocreatine kinase and 5 mg of ADP into 5 ml of
buffer. Mix by inversion.
Procedure for Assay (Note: Run everything in triplicate)
Part 1.
1) Fill three wells with a blank (10.00 µL dH2O per well)
2) a. Vortex each concentration mixture before pipetting
b. Fill the next five wells with 10.00µL of varying concentrations of ATP
standard (0.05 mM, 0.1 mM, 0.2 mM, 0.3 mM, 0.4 mM)
3) a. Vortex each concentration mixture before pipetting
b. Fill the next five wells with 10.00 µL of varying concentrations of the
PCr standard. (0.1 mM, 0.2 mM, 0.4 mM, 0.8 mM, 1.2 mM)
4) a. Vortex each sample before pipetting
b. Add 10.00 µL of sample to the appropriately wells
5) Add 185 µL of buffer to each well
6) Incubate for 25 minutes.
7) Read the plate at sensitivity of 80 (excitation setting 340, emission setting
460) (base line reading)
Part 2.
1) Add 6 µL of dilute Hexokinase to all of the wells
2) Place in the dark for 80 minutes
3) Read the plate (excitation setting 340, emission setting 460)
(R2-R1= reflects ATP in extract)
Part 3.
1) Add 6 µL of dilute Creatine Kinase to all of the wells
2) Place in the dark for 120 minutes
3) Read the plate (excitation setting 340, emission setting 460)
90 (R3-R2= reflects PCr in extract)
ATP (Sigma A-7699) Standard Curve
make fresh 5.51 mg into 5 ml dH2O
Conc (mM)
Stock (µL)
dH2O (µL)
0.05
25
975
0.1
50
950
0.2
100
900
0.3
150
850
0.4
200
800
Phosphocreatine (Sigma P-7936) Standard Curve
Stored in 7.6 mM aliquots in the -80ºC
To make 7.6 mM stock: FW (no water) 255.1 mg phosphocreatine into 50 ml
dH2O
Conc (mM)
Stock (µL)
dH2O (µL)
0. 076
10
990
0.152
20
980
0.304
40
960
0.608
80
920
0.912
120
880
C4H8N3O5PNa 4.4 mol H2Omol-1 substance
91 Therefore its effective weight is 334.3 gmol-1
II. Tyrosine Assay
1) Make up 0.1% nitroso-napthol solution (20 mg 1-nitroso-2-naphthol into
20mL methanol). Mix thoroughly.
2) Make up 2.5% sodium nitrite solution (25mg sodium nitrite into 1mL of
water). Vortex.
3) Make up 1:5 nitric acid solution (5mL nitric acid into 25mL water). Mix
thoroughly.
4) Combine 24.5mL nitric acid solution with 0.5mL of sodium nitrite solution.
Mix thoroughly.
5) Add 1 mL of nitroso-naphthol solution and 1mL of nitric acid-sodium nitrite
solution to all samples. Mix thoroughly.
6) Place samples in water bath of 65°C for 30 minutes. Allow samples to cool
post-bath.
7) After cooling, add 10mL of ethylene dichloride to each sample. Shake
samples.
8) Centrifuge samples 2000rpm at 4°C for 10 min.
9) Transfer aqueous supernatant layer of samples to a well-plate in triplicate
for fluorometric reading.
10) Open KC4. Set conditions as:
-­‐ End-point
-­‐ Fluorescence
-­‐ Read samples from both top and bottom
-­‐ Activation: 485 Emission: 590
-­‐ Sensitivity: 35 from bottom. 44 from top
11) Create a standard curve. Run fluorescence readings against the curve for
tyrosine concentrations.
III. Protein Synthesis Protocol
1) Prepare 0.61mol/L trichloroacetic acid solution (4.983g trichloroacetic acid
into 50mL of dH2O).
2) Prepare 1mol/L NaOH solution (1.999g NaOH into 50mL of dH2O).
3) During 75 minute incubation add 0.5mmol/L of L-[14C]-U-phenylalanine
(9.25MBq/L) into incubation baths with media (1.5µL phenylalanine/15mL
incubation media).
92 4) After 75 minute incubation, remove muscle from incubation bath, blot dry
and freeze in liquid nitrogen. Weigh each muscle and record.
5) Add 100µl of trichloroacetic acid solution to chilled glass homogenizer.
Additionally, add 10µl of trichloroacetic acid solution for every 1mg of
muscle. Homogenize thoroughly on ice.
6) Following homogenization, centrifuge homogenates 10,000rpm 4°C for 10
minutes.
7) Use trichloroacetic acid solution to wash the resulting pellet 3 times (100µl
each time) to eliminate non-bound radioactivity. Store supernatant for
future analysis.
8) Re-suspend pellet in 500µL of 1 mol/L NaOH and shake until solubilised.
9) Load 400µl of solubilised sample and 7.6mL of ScintiSafe solution into
scintillation tube and load into scintillation counter. Count for 14C.
10) Load 400µl of sample supernatant and 7.6mL of ScintiSafe solution into
scintillation tube and load into scintillation counter. Count for 14C.
11) Calculate protein synthesis by dividing the values incorporated in muscle
by the non-incorporated values found in the supernatant. Multiply result by
0.5mmol/L to obtain µmol of phenylalanine incorporated/mg muscle
tissue/75 minutes.
IV. Western Blotting
I.
Sample Preparation
1) Make up homogenizing buffer (250mM sucrose, 100mM KCl, 10mM
EGTA, 250mL H2O). Fix pH to 6.8 and store in fridge at 4°C.
2) Remove 10mL of homogenization buffer and add 1 tablet of protease
inhibitor (#11836170001, Roche, Laval, QC) and phosphatase inhibitor
(#04906837001, Roche, Laval, QC).
3) Weigh muscle and prepare glass homogenizers with an appropriate
amount of homogenization buffer based on the table below:
Muscle Weights
Homogenization
Buffer Added
50mg
700ul
60mg
850ul
70mg
1000ul
80mg
1150ul
93 90mg
1300ul
100mg
1450ul
110mg
1600ul
120mg
1750ul
130mg
1900ul
4) Place the glass homogenizer in ice and homogenize muscle tissue
thoroughly.
5) Once completed, collect homogenate in an eppendorf tube and store at 80°C.
II.
Protein Determination
1) Remove BSA standard (1g/mL) from -20°C freezer and thaw.
2) Make up standards based on table below:
Final Protein
Volume of BSA
Concentration (mM)
(µ
µ L)
0
0
1000
0.05
100 from '0.5'
900
0.125
500 from '0.25'
500
0.25
500 from '0.5'
500
0.5
500 from '1'
500
1
1000
0
94 Volume of dH2O (µ
µ L)
3) Pipette 10µl of each standard concentration in triplicate in a clear
microtitre plate. Similarly, pipette all samples in triplicate as 1µl and 9µl
H2O.
4) Add 200µl of diluted 1:4 Bio-Rad Protein Assay Dye Reagent (#5000006,
BioRad, Mississauga, ON) to all wells and let incubate for 5 min. Remove
all air bubbles.
5) Open KC4 software. Read absorbance at 595nm. Intensity: 3, Duration: 5
6) In Excel, run a polynomial curve with the blank subtracted from all wells.
Use the curve equation to determine the y-intercept and multiply by 10 to
determine the protein concentration of each sample.
III.
SDS PAGE
1) To prepare samples for loading, follow table below:
Total Volume (TV)
Sample Volume (SV)
[Desired Final Protein]
[Protein]
TV
TV
(i.e.
for 5x SB i )
SB f =
[ SBi ]
5
dH 2O = TV − SV − SB
SV = TV Sample (Laemelli) Buffer (SB)
Water (H2O)
**Desired final protein concentration is 1µg/µl**
2) Boil samples for 5 minutes and subsequently place on ice for 5 min.
3) Clean all glass plates with methanol and place in glass holding apparatus
with small plate facing inward. Ensure glass plates are flush before
inserting the apparatus into the gel casting stand.
4) To prepare gels, follow chart below:
95 Protogel (mL)
Gel Buffer (mL)
Percent Gel
dH2O (mL)
4 (Stacking Gel)
6.1
1.3
2.5 stacking
4.85
2.5
2.5 resolving
10
4.01
3.33
2.5 resolving
12
3.33
4.01
2.5 resolving
7.5 (Running
Gel)
(30% acrylamide)
5) Before pouring running gel, add 100µl of Ammonium Persulfate (APS)
(60mg/600µl) and 10µl of Tetramethylethylenediamine (TEMED). Swirl
and pour until reaching the depth of the combs. Remove any bubbles with
methanol and let gel harden.
6) Once running gel has hardened, pour off methanol. Before pouring
stacking gel, add 100µl of APS and 20µl TEMED. Swirl and pour to the top
of small glass plate and place comb in. Once hardened, remove comb and
rinse wells with dH2O.
7) Place gel cassettes into electrode assembly with the short plates facing
inwards, clamp down firmly and place into mini-tank.
8) Prepare running buffer (250mM Tris Base, 1.92M glycine, 1% SDS, 1L
dH2O, pH to 8.3. Take 50mL of stock solution running buffer and add
450mL dH2O). Fill inner chamber with running buffer and add
approximately 250mL to outside chamber.
9) Load 5µl of molecular weight standard (#161-0375, BioRad, Mississauga,
ON) into one well of each gel and an appropriate amount of sample into
each well of the gel (for protein specific information, see Protein Specific
Western Blotting Information Table).
10) Run electrophoresis specific to the protein of interest (for protein specific
information, see Protein Specific Western Blotting Information Table)
IV.
Transfer
96 1) Prepare fresh transfer buffer (312.5mM Tris Base, 2.4M glycine, 100mL
dH2O; Take 80mL of solution, add 200mL methanol and 720mL dH2O).
2) Equilibrate sponge pads, fibre pads and filter paper in transfer buffer in a
plastic tray for 10 minutes. For PVDF membrane, pre-soak in methanol for
10 seconds, then equilibrate in transfer buffer. For nitrocellulose
membrane, pre-soak in dH2O for 10 seconds, then equilibrate in transfer
buffer.
3) Remove gels from glass plates, discard stacking gel and equilibrate gel in
transfer buffer for approximately 2 minutes.
4) Create transfer sandwiches: clear side of transfer cassette, sponge pad,
fibre pad, 2 filter paper, membrane (PVDF or nitrocellulose), gel, 2 filter
paper, fibre pad, sponge pad, black side of transfer cassette. Roll out any
air bubbles in sandwich.
5) Fill tank with transfer buffer and place tank in an ice bath. Add stir bar into
the tank to help dissipate heat. Place cassettes into transfer case with the
black side of the cassette to the black side of the case. Set voltage and
transfer time according to protein of interest (for protein specific
information, see Protein Specific Western Blotting Information Table).
V.
Antibodies
1) Discard filter papers and gel. Transfer membranes to petri dish. Block
membranes with milk-TBST solution (20mM tris base, 137mM NaCl, and
0.1% (v/v) tween 20, pH 7.5) solution. % of milk and blocking time
dependent on protein of interest (for protein specific information, see
Protein Specific Western Blotting Information Table).
2) After appropriate amount of time discard blocking solution and add 1°
antibody solution to membrane in the appropriate dilution overnight (for
protein specific information, see Protein Specific Western Blotting
Information Table).
3) After overnight incubation, discard 1° antibody solution and wash the
membrane 4 times for 5 minutes each time in TBST solution.
4) After 4 washes, add 2° antibody solution to the membrane in the
appropriate dilution. Let rock at room temperature. 2° antibody, solution,
dilution, and time all dependent on protein of interest (for protein specific
information, see Protein Specific Western Blotting Information Table).
VI.
Enhanced Chemi-Luminescence
97 1) After appropriate amount of time discard the 2° antibody solution and
wash the membrane again 4 times for 5 minutes each time in TBST
solution.
2) After 4 washes, combine 1mL each of Chemi-Luminescent HRP Substrate
Peroxide Solution and Luminal Reagent (#WBKLS0500, Billerica, MA)
with the membrane and pipette over for 5 minutes.
3) Blot membrane dry and place between two transparency sheets. Roll out
any air bubbles.
4) Expose the membrane using the FluroChem imaging system (AlphaInnotec, Santa Clara, CA), and quantify using the AlphaEaseFC Software
(Alpha-Innotec, Santa Clara, CA). Exposure time dependent on protein of
interest (for protein specific information, see Protein Specific Western
Blotting Information Table). Determine the density of the bands using
“Image J” software (National Institutes of Health, Bethesda, MD).
98 Protein Specific Western Blotting Information
mTOR
Phosph
o-mTOR
p70s6k
Phosphop70s6k
Ubiquitin
Conjugates
Actin
µ Calpain
Protein
Load
15µl
15µl
10µl
10µl
12µl
12µl
10µl
Stacking
Gel
4%
4%
4%
4%
4%
4%
4%
Running
Gel
7.5%
7.5%
10%
10%
7.5%
7.5%
7.5%
Gel
Electroph
oresis
2 hours
at 120V
2 hours
at 120V
Membrane
PVDF
PVDF
2 hours
at 100V
2 hours
at 100V
90 min at
100V
90 min at
100V
75 min at
100V
75 min at
100V
Nitrocellulose
90 min at
100V
1% milkTBST for
2 hours
1% milkTBST for
2 hours
2.5% milkTBST for 1
hour
2.5% milkTBST for 1
hour
5% milkTBST for 1
hour
5% milkTBST for 1
hour
5% milkTBST for 1
hour
1:1000
rabbit
polyclona
lin
1%milkTBST
1:7000 in
1% milkTBST
antirabbit for
2 hours
1:1000
rabbit
monoclo
nalin 1%
milkTBST
1:7000 in
1% milkTBST
antirabbit for
2 hours
1:1000
rabbit
polyclonal
in 5%
milk-TBST
1:1000
goat
monoclon
al in 5%
milk-TBST
1:2000 rabbit
polyclonal in
5% milkTBST
1:5000
mouse
monoclon
al in 5%
milk-TBST
1:6000
anti-rabbit
in 5%
BSATBST for 1
hour
1:6000
anti-goat
in 5%
BSATBST for 1
hour
1:6000 antirabbit in 5%
milk-TBST
for 1 hour
1:5000
antimouse in
5% milkTBST for 1
hour
1:1000
rabbit
monoclon
al in 1%
BSAPBST
1:5000
anti-rabbit
in 5%
milk-TBST
for 1 hour
2 min
2 min
2 min
2 min
2 min
2 min
Transfer
Blocking
1°
2°
Exposure
90 min at
120V
PVDF
90 min at
120V
PVDF
99 75 min at
100V
PVDF
75 min at
100V
PVDF
15 min at
100V
followed
by 60 min
at 160V
Cut
membrane
at 43kDa.
30 min
V. Western Blotting Images
1) Total mTOR
100 2) Phospho-mTOR
101 3) Total p70s6 kinase
102 4) Phospho-p70s6 kinase
103 5) Ubiquitin Conjugates and Actin
104 6) µ-Calpain
105