California State University, Northridge

CALIFORNIA STATE UNIVERSITY, NORTHRIDGE

Effects of Silencing Lipoprotein Lipase on Metabolic Enzymes in Rat Muscle Cells

A thesis submitted in partial fulfillment of the requirements

For the degree of Master of Science in Biochemistry

Adam Scott Mogul

May 2019

The thesis of Adam Mogul is approved:

______Dr. Simon Garrett Date

______Dr. Daniel Tamae Date

______Dr. Jheem Medh, Chair Date

California State University, Northridge

Acknowledgements

I would like to thank everyone in the CSUN Chemistry and Biochemistry department for helping me along this journey, especially those of you that helped supply me with helpful tips, reagents, or buffer solutions in my times of dire need. All of my friends in the department, the graduate and undergraduate students that have made my time at CSUN so enjoyable. The great group of people that have been a part of Dr.

Medh’s lab, both past and present. Finally, a heartfelt thank you to Dr. Medh, for all of the help, understanding, and support through the time that I have spent in this program.

Your guidance and patience have provided the foundation for my successful navigation of the world of biochemistry, and I truly appreciate it.

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Dedication

I would like to dedicate this thesis to my Mom and Dad, who have believed in me and supported me through my academic career and beyond. To my grandparents, who have genuinely tried to be interested when I talk about my research, and provided me with love, food, and a place to sleep near school on late nights. To my sister, for being my life-long companion. And to my friends, new and old, who understood when I was too busy to see them but still made the effort to see me when I was available.

Table of Contents

Signature Page ii

Acknowledgements iii

Dedication iv

List of Tables viii

List of Figures ix

List of Abbreviations xi

Abstract xiii

Chapter 1 – Introduction

Section 1 – Prevalence and mechanism of diabetes 1

Section 2 – Muscle cells and metabolism 1

Section 3 – Lipoprotein Lipase (LPL) 2

Section 4 – Physiological role of LPL products 6

Section 5 – Glycolysis, glycogen synthesis, and LPL 6

Section 6 – Previous study of LPL and its impact on

glucose utilization 10

Section 7 – Research goals 10

Chapter 2 – Materials and Methods

Section 1 – L6 rat muscle cell culture 12

Section 2 – L6 rat muscle cell differentiation 13

Section 3 – The LPL-knock-down L6 cell line 14

Section 4 – RNA isolation 15

Section 5 – RNA quantification 16

Section 6 – RT-PCR to make cDNA 17

Section 7 – End-point PCR 18

Section 8 – Quantitative PCR 20

Section 9 – L6 cell lysis for protein isolation 22

Section 10 – Protein quantification assay 22

Section 11 – Western blot 24

Section 12 – Hexokinase activity assay 27

Section 13 – Statistical analysis 32

Chapter 3 – Results

Section 1 – Morphology of LPL-KD cells 33

Section 2 – Effects of LPL-KD on RNA expression levels 34

Section 3 – Effects of LPL-KD on protein expression levels 36

Section 4 – Effects of LPL-KD on hexokinase activity 37

Chapter 4 – Discussion

Section 1 – Effect of LPL on gene expression 45

Section 2 – Effect of LPL on hexokinase II protein levels 47

Section 3 – Effect of LPL on the activity of hexokinase 49

Section 4 – Literature review of altered LPL expression

and metabolism 55

Chapter 5 – Conclusion 58

References 60

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List of Tables

Table 1. Primers used for PCR reactions. 19

Table 2. Antibodies used in western blotting 26

Table 3. Baseline NADH formation in HK assay samples. 41

Table 4. Raw absorbance value comparison of concentrated lysates. 42

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List of Figures

Figure 1. Overview of the processing and transport of dietary lipids. 3

Figure 2. LPL catalyzes the hydrolysis of triglyceride into glycerol and

free fatty acids 3

Figure 3. LPL dimer bound to GPIHBP1. 4

Figure 4. The catalytic triad of the active site of LPL. 5

Figure 5. Hexokinase catalyzes the phosphorylation of glucose to

glucose 6-phosphate 7

Figure 6. Phosphorylation of fructose 6-phosphate to

fructose 1,6-bisphosphate by the enzyme phosphofructokinase-1. 9

Figure 7. Parallel differentiation of wild-type cells for experimentation. 14

Figure 8. Theory behind the hexokinase activity assay. 28

Figure 9. Comparison of the morphology of the L6 WT vs LPL-KD cells. 33

Figure 10. End-point PCR products separated via 2% agarose gel electrophoresis. 34

Figure 11. Regulation of gene transcription in LPL-KD cells as determined

using qPCR. 35

Figure 12. Western blot analysis of HK II protein concentration in L6 cells. 37

Figure 13. Standard curve used for the hexokinase assay data analysis. 38

Figure 14. Hexokinase activity in WT and LPL-KD L6 cells. 39

Figure 15. Comparison of the nmol of NADH formed per mg of protein

per minute for each sample. 40

Figure 16. Comparison of NADH formation in the concentrated WT

and KD samples after background-subtraction. 43

Figure 17. Summary of the main findings of this research. 44

Figure 18. Proposed explanation for the regulation of hexokinase II activity

in LPL-KD cells. 58

List of abbreviations

6PG 6-phospho-D-glucono-1,5-lactone APS Ammonium persulfate BCA Bicinchoninic acid Bis-acrylamide N’N’-bis-methylene-acrylamide BG sample Background sample BSA Bovine serum albumin cDNA Complementary DNA

CT value Cycles required to reach fluorescence threshold DMSO Dimethyl sulfoxide dsDNA Double-stranded DNA F16-BP Fructose 1,6-bisphosphate F6-P Fructose 6-phosphate faCoAs Fatty acyl-CoAs FBS Fetal bovine serum FFA Free fatty acid G6-P Glucose 6-phosphate G6PD Glucose 6-phosphate dehydrogenase GPIHBP1 Glycosylphosphatidylinositol-anchored high-density lipoprotein binding protein 1 HK Hexokinase hLPL0 Heart-specific LPL knock-out mice HRP Horseradish peroxidase IDL Intermediate-density lipoprotein IRS-1 Insulin receptor substrate-1 KD Knock-down (of LPL unless otherwise noted) LCACoA Long-chain acyl CoA LPL Lipoprotein lipase

LPL-KD Lipoprotein lipase-knock-down M-MLV Moloney Murine Leukemia Virus M-MLV MM Moloney Murine Leukemia Virus master mix M-MLV RT Moloney Murine Leukemia Virus Reverse Transcriptase mRNA Messenger RNA PBS Phosphate buffered saline PCR Polymerase chain reaction PFK Phosphofructokinase PI Phosphatidylinositol PMSF Phenylmethanesulfonyl fluoride PPP Pentose phosphate pathway qPCR Quantitative PCR rcf Relative centrifugal force RNAi RNA interference RT-PCR Reverse transcriptase polymerase chain reaction SDS Sodium dodecyl sulfate SDS-PAGE Sodium dodecyl sulfate – polyacrylamide gel electrophoresis shRNA Short hairpin RNA T1DM Type 1 diabetes mellitus T2DM Type 2 diabetes mellitus TEMED N,N,N’,N’-tetramethylethylenediamine TG Triglyceride VLDL Very-low-density lipoprotein WT Wild-type

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Abstract

Effects of Silencing Lipoprotein Lipase on Metabolic Enzymes in Rat Muscle Cells

Adam Scott Mogul

Master of Science in Biochemistry

Lipoprotein lipase (LPL) is an enzyme required for the hydrolysis of triglycerides to free fatty acids and glycerol. In earlier reports, we demonstrated that in a muscle cell line, LPL levels are directly correlated with insulin resistance, and reducing LPL expression increased insulin-stimulated glucose uptake. Additionally, LPL-knock-down

(LPL-KD) L6 rat muscle cells showed increased glucose oxidation compared to wild- type (WT) L6 cells. Glycolysis is the first pathway in glucose oxidation, and two important enzymes for regulating glycolysis in skeletal muscle are hexokinase II (HK II) and phosphofructokinase-1 (PFK-1). Our goal was to compare the expression and activity of the glycolytic enzymes hexokinase and phosphofructokinase in LPL-KD and

WT L6 muscle cells.

The cells used in this project were prepared in a previous study. For LPL-KD cells, the LPL gene was silenced in rat skeletal muscle cells (L6 cells) by lentiviral- mediated RNA interference. Total RNA was isolated and specific primer pairs were used

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for end-point and real-time PCR amplification of LPL, Hexokinase II, and PFKM. A western blot analysis was carried out for HK II protein mass. β-actin was used as a housekeeping gene during all PCR and western blot analyses. Additionally, HK activity assays were performed to determine the HK enzyme activity in the cell lysates.

Quantitative PCR showed that LPL-KD cells have <1% of the LPL expression as their WT counterparts due to the shRNA silencing. Silencing of the LPL gene resulted in a dramatic increase in the gene transcription of HKII (253% of WT levels) and a decrease in that of PFKM (72.1% of WT levels). These findings were supported by data from end- point PCR and western blot analysis of hexokinase II. The hexokinase activity assay suggested a decrease in HK activity in LPL-KD cells compared to WT cells.

We hypothesize that increased HK II expression and translation facilitates glucose uptake into the cell. The finding that the glycolytic rate-limiting enzyme PFK-1 is repressed suggests that glucose 6-phosphate may be diverted to glycogen synthesis, the pentose phosphate pathway, or generally build up to a high enough concentration to inhibit the overall HK II activity, in spite of the increased amount of HK II protein in the

LPL-KD cells.

xiv Chapter 1 – Introduction Section 1 – Prevalence and mechanism of diabetes

Diabetes mellitus is a widespread and chronic metabolic disease that afflicts an estimated 8.3% of the world’s adult population.1 The disease can be categorized into two main types and is characterized by hyperglycemia, which leads to complications that include cardiovascular diseases, neuropathy, retinopathy, and nephropathy. Type 1 diabetes mellitus (T1DM) is caused by an autoimmune response against the β (beta) cells of the pancreas, which are responsible for the secretion of insulin. Destruction of the β cells leads to the inability of a patient with T1DM to produce enough insulin to regulate metabolic needs, which results in hyperglycemia.

Type 2 diabetes mellitus (T2DM) is the most common type of diabetes, accounting for approximately 87-91% of all diabetes cases.2 In patients with T2DM, the tissues targeted by insulin develop insulin resistance. As a result, more insulin is required to maintain the desired blood glucose levels, and hyperglycemia often occurs despite the production of insulin by pancreatic β cells. One of the primary targets of insulin activity is skeletal muscle, which plays an important role in maintaining glucose homeostasis.

Section 2 – Muscle cells and metabolism

Skeletal muscle is one of the major tissues involved in insulin-mediated glucose uptake, and is responsible for as much as 38% of whole body glucose uptake.3 The glucose taken in by the muscle can be used to produce ATP via glycolysis in times of light exercise or rest. Alternatively, the glucose can be stored as glycogen in the muscle cells. This glycogen can later undergo lactic acid fermentation in order to replenish ATP

in the muscles during bursts of heavy activity.4 However, under conditions of low to moderate exercise, the muscle cells primarily utilize fatty acids or ketone bodies as their fuel sources.4 As such, it is critical that muscle cells are able to obtain both glucose and fatty acids from the bloodstream.

Section 3 – Lipoprotein lipase (LPL)

Lipids, such as fatty acids and cholesterol, are packaged into lipoproteins to facilitate transport through the blood plasma (Figure 1). Chylomicrons and very-low- density lipoproteins (VLDLs) are the types of lipoproteins that are of particular importance to metabolism, as they contain a high weight percent of triglycerides (TG).4

These triglycerides can be utilized by the enzyme lipoprotein lipase in order to produce fatty acids that are required for the energy production in muscles.

Lipoprotein lipase (LPL) catalyzes the hydrolytic cleavage of TGs found in chylomicrons and VLDLs into free fatty acids (FFA), monoglycerides, glycerol, and intermediate-density lipoproteins (IDLs).5 This hydrolysis is summarized in Figure 2.

LPL is a secreted enzyme that is produced in myocytes, adipocytes, and macrophages.6

The secreted LPL is localized to the luminal side of the capillary endothelium through the transport and anchoring actions of glycosylphosphatidylinositol-anchored high-density lipoprotein binding protein 1 (GPIHBP1).7 The LPL remains bound to the GPIHBP1 while it is active in the capillaries.8 LPL activation depends on apolipoprotein C-II, which is found in chylomicrons and VLDLs.5

Figure 1. Overview of the processing and transport of dietary lipids. 1) Bile salts in the small intestine emulsify dietary fats to form mixed micelles. 2) Intestinal lipases degrade the triglycerides from the mixed micelles. 3) Lipase products, including fatty acids, are taken up by the intestinal mucosa, where they are converted back to triglycerides. 4) Triglycerides, cholesterol, and apolipoproteins are incorporated into chylomicrons. 5) Chylomicrons move through the bloodstream to the tissues. 6) Lipoprotein lipase in the capillaries converts triglycerides into fatty acids and glycerol. 7) The fatty acids enter the cells, which are either myocytes or adipocytes. 8) Fatty acids are oxidized as fuel in the muscle or re-esterified for storage in adipocytes. Figure adapted from Lehninger Principles of Biochemistry, 5th edition.4

Figure 2. LPL catalyzes the hydrolysis of triglyceride into glycerol and free fatty acids. The R groups are hydrocarbon chains of various lengths and degrees of saturation.

The active form of the LPL enzyme is a homodimer that assembles in a head-to- tail fashion.9 The substrate recognition site on one subunit facilitates the substrate interaction that allows the catalytic site on the other subunit to carry out the hydrolysis reaction, and vice versa. The two subunits interact in a non-covalent manner at the dimer interface.10 The crystal structure of LPL in complex with GPIHBP1 was recently solved by Birrane, et al, and is shown in Figure 3.

Figure 3. LPL dimer bound to GPIHBP1. The GPIHBP1 is shown in blue. The second LPL subunit was shown as a solid green color in cartoon form on the left, then in surface form on the right in order to highlight the dimer interactions. Each LPL subunit has an N-terminal α/β-hydrolase domain, comprised of 6 α- helices and 10 β-strands, which contains the catalytic triad of serine 159, aspartate 183, and histidine 268 (Figure 4).8 The C-terminus has a flattened β-barrel domain that is comprised of 12 β-strands. The two termini are connected by a hinge region. LPL also contains multiple, positively charged heparin binding domains,8 which enable the enzyme to bind to extracellular heparan sulfate proteoglycans on muscle cells until the GPIHBP1

binds and localizes the enzyme to the capillaries.11,12 The catalytic sites of both subunits are on the same side of the dimer and in the same plane, which suggests that both subunits are active on the same substrate (chylomicron or VLDL) at the same time.8–10

Figure 4. The catalytic triad of the active site of LPL. The residues serine 159, aspartate 183, and histidine 268 are critically important for the activity of the hydrolase domain. The binding of GPIHBP1 to LPL, which is non-covalent and mostly due to hydrophobic interactions, is critically important to LPL-mediated lipid metabolism.8

GPIHBP1 facilitates the proper localization of LPL to the capillaries, but it also stabilizes the structure of LPL. The catalytic hydrolase domain of LPL is susceptible to both spontaneous unfolding as well as unfolding by physiological inhibitors of LPL, such as

ANGPTL4 and ANGPTL3.8 However, when LPL is in complex with GPIHBP1, the frequency of unfolding is dramatically reduced, thus the catalytic activity is preserved.13

Additionally, GPIHBP1 plays a role in sequestering the triglyceride-rich lipoproteins to the walls of the capillaries from the bloodstream so that LPL is able to perform its catalysis.14

Section 4 – Physiological role of LPL products

As previously mentioned, LPL hydrolyzes triglycerides into three FFAs and glycerol. However, since monoglycerides have the potential to diffuse across cell membranes, the physiological end result of the lipolysis can be a monoglyceride and two

FFAs.4,5 These products of lipoprotein lipase activity, particularly free fatty acids, have important roles in human physiology.

Circulating and dietary triglycerides provide more than half of the energy required for resting skeletal muscle, the liver, and the heart.4 For skeletal muscle, LPL hydrolysis of the TGs in chylomicrons or VLDLs is the first step to obtaining this energy source.

The FFAs that enter the muscle cells as a result of this hydrolysis are then activated by the carnitine shuttle in order to be transported across the mitochondrial membrane for oxidation.4,15 The first reaction of the carnitine shuttle converts FFAs into fatty acyl-

CoAs (faCoAs). These fatty acyl-CoAs can either continue along the carnitine shuttle to be oxidized for energy in the mitochondria, or can remain in the cytosol for use in the synthesis of membrane lipids.4 However, there is evidence that cytosolic accumulation of faCoAs can lead to impaired insulin signaling and insulin resistance.16

Section 5 – Glycolysis, glycogen synthesis, and LPL

Glycolysis is the tightly regulated metabolic pathway in which a molecule of glucose is degraded into two molecules of pyruvate, yielding a net production of 2 molecules of ATP. It is the first stage of the complete breakdown of glucose molecules for energy, and is followed by the citric acid cycle under aerobic conditions, or lactic acid fermentation under anaerobic conditions.4 The first five enzyme-catalyzed reactions of

glycolysis are referred to as the preparatory phase. Two of those reactions, catalyzed by the enzymes hexokinase (HK) and phosphofructokinase (PFK) consume energy in the form of ATP in order to produce higher energy metabolic intermediates. These energy- consuming reactions are irreversible, and subject to regulation at multiple levels.

Hexokinase

In the first step of glycolysis (Figure 5), hexokinase catalyzes the irreversible,

ATP-dependent phosphorylation of glucose into glucose 6-phosphate (G6-P).4 The primary isozyme of HK found in muscle cells, hexokinase II (HK II), has the highest affinity for glucose of all HK isozymes.17 In fact, the glucose concentration in muscle cells is generally high enough to saturate the enzyme, causing it to perform catalysis at nearly its maximal rate.4 However, HK II in muscles is reversibly and allosterically inhibited by its product, G6-P. Thus, when G6-P levels are elevated, HK II activity is reduced, which helps to reestablish the balance between the production and utilization of

G6-P.

Figure 5. Hexokinase catalyzes the phosphorylation of glucose to glucose 6-phosphate.

The glucose 6-phosphate that is produced by the reaction catalyzed by hexokinase does not necessarily commit the glucose molecule to the glycolytic pathway. There are several other fates that can befall the G6-P, including oxidation to pentose phosphates via the pentose phosphate pathway, or use as the starting point for the synthesis of glycogen.4

The pentose phosphate pathway (PPP) plays a fairly small role in healthy skeletal muscle, but it does occur to some extent under anaerobic conditions.18 Glycogen synthesis is more relevant to skeletal muscles, as glycogen is used as one of the sources of ATP during times of heavy exertion.4 However, glycogen only makes up about 1% of the total weight of skeletal muscle, so most of the G6-P produced in the muscle is used for glycolysis.

Hexokinase inhibition does not only occur as a result of a buildup of the G6-P product. There are other manners of regulating the activity of HK II, including allosteric inhibition by other molecules, and transcriptional regulation.4 One such class of allosteric inhibitors are the fatty acyl-CoAs,19 specifically palmitoyl CoA (16:0), oleoyl CoA (18:1, n=9), and linoleoyl CoA (18:2, n=6).20 These three faCoAs, the three major fatty acyl-

CoA species that are found in skeletal muscle, were shown to inhibit hexokinase activity in a concentration-dependent manner while glucose was present in physiological concentrations. This inhibition of HK activity by these faCoAs provides a link between the uptake of the LPL products by a muscle cell and the ability of that cell to start glucose metabolism. Additionally, the reduced capability of the muscle cell to perform glycolysis or utilize glucose may also result in reduced insulin sensitivity of that cell.20

Phosphofructokinase

In the third overall step of glycolysis, the enzyme phosphofructokinase-1 (PFK-1) catalyzes the ATP-dependent phosphorylation of fructose 6-phosphate (F6-P) to fructose

1,6-bisphosphate (F16-BP). This reaction, summarized in Figure 6, is the second irreversible step of glycolysis that consumes ATP, but the first “committed” step of glycolysis.4 As mentioned previously, the G6-P that is formed as a result of HK activity is able to be utilized in multiple pathways. However, the F16-BP produced as a result of the PFK-1 activity is only used in glycolysis. As a result, PFK-1 is the subject of complex allosteric and transcriptional regulation by the cell.4

Figure 6. Phosphorylation of fructose 6-phosphate to fructose 1,6-bisphosphate by the enzyme phosphofructokinase-1. Phosphofructokinase-1 in muscle cells is a homotetrameric protein, made from four identical subunits that are encoded by the PFKM gene.21 PFK-1 utilizes ATP as a substrate, but ATP is also produced as the result of glycolysis. When it is at a sufficiently high concentration, ATP is able to bind to an allosteric site and lower the affinity of PFK-

1 for the substrate F6-P, thus reducing the enzyme’s activity.4 In contrast, when the concentrations of ADP and AMP, molecules that result when more ATP is being used than being formed, reach sufficiently high levels, they act to allosterically relieve the inhibition by ATP and restore the activity of the PFK-1.

Reversible regulation of PFK-1 by fatty acyl-CoAs has also been shown to occur in muscle cells.22 This inhibition occurs at physiological concentrations of the faCoAs, but is prevented by sufficient concentrations of ADP or AMP. The presence of ATP was not demonstrated to prevent this inhibitory effect. These findings further provide evidence that the uptake of free fatty acids into the cell causes reduced utilization of glucose for energy production, which may contribute to insulin resistance.

Section 6 – Previous study of LPL and its impact on glucose utilization

Previous studies in our research group have focused on measuring the effects of silencing LPL on glucose uptake and insulin sensitivity in rat skeletal muscle cells (L6 cells). It has been shown that the downregulation of LPL expression leads to increased glucose uptake by the muscle cells, both in the absence and presence of insulin.23

Additionally, it was demonstrated that glycogen synthesis and glucose oxidation in LPL- knock-down (LPL-KD) L6 cells were slightly higher than those of the wild-type (WT) L6 cells in the absence of insulin, but approximately twice as high as the WT in the presence of insulin.24 These findings confirm that LPL and its lipolysis products reduce insulin sensitivity in muscle cells.

Section 7 – Research goals

In an effort to further explore the relationship between lipoprotein lipase and the glycolytic pathway, we wanted to compare WT L6 cells with LPL-KD L6 cells in order to determine:

1. Is there a difference in gene expression of the glycolytic enzymes hexokinase II and phosphofructokinase-1 between the two cell lines? It has been proven that fatty acyl-

CoAs reduce the activity of the enzymes, but there appears to be far less research into the long-term transcriptional effects that result from the absence or reduction of LPL during the muscle cell life cycle.

2. If there is a difference in gene transcription, is that difference carried over to the protein level? The central dogma of biology states that DNA is used to produce

RNA, which in turn is used as a template for proteins. Therefore, if there is a difference observed in the gene transcription levels, then having protein data that follows the same trend would support those findings.

3. Is there a difference in the activity level of the glycolytic enzymes between the

WT L6 cells and the LPL-KD L6 cells? Gene expression and protein levels are not necessarily correlated with enzyme activity, so it would be interesting to see if there is any change in activity observed in enzymes from the different cell lines.

Chapter 2 – Materials and Methods

Section 1 – L6 rat muscle cell culture

L6 cells are skeletal muscle cells isolated from the Rattus norvegicus rat, and were purchased as myoblasts from American Type Culture Collection (ATCC, Rockville,

MD). The cells were received frozen, in freezing medium made of 95% growth medium with 5% dimethyl sulfoxide (DMSO). Growth medium is 10% fetal bovine serum (FBS) and 90% complete DMEM, which is Dulbecco’s Modification of Eagle’s Medium

(DMEM) with 4.5 g/L glucose, L-glutamine & sodium pyruvate from Corning cellgro

(Ref # 10-013-CM), supplemented with an additional 50 units/mL penicillin, 50 µg/mL streptomycin, 10 mM HEPES, and 2 mM glutamine.

The frozen cells were rapidly thawed in a 37 ºC water bath, sterile transferred to a

15-mL polystyrene conical tube (Falcon) in a laminar flow tissue culture hood (NUAIRE

Biological Safety Cabinet, Class II Type A/B3), and diluted with 11 mL of sterile-filtered phosphate buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.4, filtered through a 0.22 µm filter). The cell suspension was then centrifuged at 800 rpm for 5 minutes in a swinging bucket centrifuge (IEC Centra GP8R) to produce a cell pellet. The supernatant was aspirated, and the cells were re-suspended in 10 mL of growth medium. The re-suspended cells were transferred to a 75 cm2 polystyrene cell culture flask, which was then placed in a humidified incubator kept at 37

ºC and 5% CO2 to facilitate cell proliferation.

The L6 cells are adherent cells, and were incubated until they reached ~80% confluency. To subculture the cells for further propagation, differentiation, or freezing,

the growth medium was aspirated from the flask. Next, the adhered cells were washed with 10 mL of sterile PBS, and 0.5 mL of 1X Trypsin/EDTA (0.05% Trypsin, 0.53 mM

EDTA from Corning cellgro) was added to the flask. The flasks were rocked briefly to ensure that the trypsin covered all of the cells, then the excess trypsin/EDTA was aspirated off and the flasks were incubated at 37 ºC at 5% CO2 for 5 minutes. Following incubation, 10 mL of PBS was added to re-suspend the cells, and this cell suspension was transferred to a 15-mL conical Falcon tube for centrifugation at 800 rpm for 5 minutes.

The supernatant was aspirated, and the cells in the pellet were re-suspended in growth medium (for differentiation or propagation) or in L6 freezing medium (95% growth medium, 5% DMSO).

Cells that were re-suspended in L6 freezing medium were sterile transferred to a cryovial for freezing. These cells were initially frozen in the vapor phase of liquid nitrogen, and then subsequently moved to the liquid phase for longer-term storage.

Section 2 – L6 rat muscle cell differentiation

In order to differentiate the L6 cells from myoblasts into mature myocytes, a portion of the cells, grown and re-suspended as discussed previously, were aliquoted into

5 mL of growth medium and transferred to a 25 cm2 polystyrene cell culture flask. The cells were left in growth medium (10% FBS, 90% complete DMEM) overnight to allow cell adhesion. The growth medium was aspirated, and 5 mL of differentiation medium 1

(2% FBS, 98% complete DMEM) was added to the cells and allowed to incubate in the same conditions as propagating cells for 2 days. Subsequently, the differentiation medium 1 was aspirated and replaced with differentiation medium 2 (0% FBS, 100% complete DMEM) for the final 2 days of incubation, after which differentiation was

considered complete. Differentiation was confirmed under the microscope by observing that the formerly oval-shaped myoblasts had elongated and aligned with surrounding cells, which would suggest differentiation into myocytes. Figure 7 shows a summary of the parallel differentiation and propagation of the cells.

Figure 7. Parallel differentiation of wild-type cells for experimentation. The smaller, numbered flasks are those which are used for differentiation. Flasks in a box are designated for the same type of experiment. For example, flasks 1 and 2 may be for protein isolation, while flask 3 is for RNA isolation. However, all 3 of the flasks come from the same cell stock, and thus should be nearly identical. The same method was used for LPL-KD cells. Section 3 – The LPL-knock-down L6 cell line

The LPL-KD L6 cell line was generated through the use of lentiviral-mediated

RNA interference (RNAi) by Majib Jan24. The lentiviral vector that was transfected into the cells included a short hairpin RNA (shRNA) transgene that targeted the LPL in the host genome of the L6 cells, as well as a puromycin resistance gene. The shRNA was responsible for knocking down the expression of LPL in the cells that underwent

transfection, while the puromycin resistance gene was included in order to allow for selection of successfully transfected cells.

The RNAi process created a stable line of LPL-KD L6 cells, which would pass the shRNA, and in turn the decreased LPL expression, down to their progeny during propagation. However, due to the potential for transfected cells to revert back to their wild-type state, the LPL-KD cells would periodically be treated with selection medium

(L6 growth medium with 10 µg/mL puromycin) for 5-7 days during propagation.

Treatment with the antibiotic puromycin had little to no effect on the LPL-KD cells that were expressing the resistance gene, but would kill any WT L6 cells or those that no longer have resistance.

LPL-KD cells were otherwise propagated and differentiated under the same conditions as described previously for the WT L6 cells. The knock-down of LPL expression was confirmed using gene-specific primers and the polymerase chain reaction, as discussed in subsequent sections.

Section 4 – RNA isolation

Following differentiation, the medium was aspirated, and the cells were washed with 5 mL of refrigerated, sterile PBS. In order to lyse the cells for RNA extraction, 1 mL of TRI Reagent (Thermo Fisher or Molecular Research Center, Inc.) was added to the washed cells in the 25 cm2 polystyrene flask. TRI Reagent, a monophase solution that contains phenol and guanidine thiocyanate, is able to lyse cells and dissolve the RNA,

DNA, and proteins that they contain. The flask containing the cells in TRI Reagent was

then placed in a -20 ºC freezer for at least 12 hours in order to assist in cell lysis and improve RNA yield.

Following the freezing, the flask of cells was removed from the freezer and allowed to equilibrate to room temperature for 10 minutes. The TRI Reagent/cell lysate solution was then transferred by micropipette (Gilson pipetman or Eppendorf Research

PhysioCare) to a 1.5-mL microcentrifuge tube (Eppendorf) under non-sterile conditions.

Next, 100 µL of chloroform was added to the solution, mixed well by inversion, and allowed to incubate at room temperature for 10 minutes. Phase separation was achieved by centrifuging the mixture at 12,000 rcf (relative centrifugal force) for 15 minutes in an

Eppendorf Centrifuge 5424. The RNA is contained in the upper, aqueous phase, which was carefully transferred to a fresh microcentrifuge tube. To precipitate the RNA out of solution, 500 µL of isopropanol was added to this aqueous phase, the solution was briefly vortexed, and then allowed to incubate at room temperature for 10 minutes.

Subsequently, the solution was centrifuged at 12,000 rcf for 8 minutes, and the supernatant was decanted off to leave behind the RNA pellet. The pellet was washed by adding 1 mL of 75% ethanol and then centrifuging at 7,500 rcf for 5 minutes. The ethanol was then decanted off, and the RNA pellet was allowed to air dry for 5-7 minutes.

To bring the RNA back into solution, 50 µL of nuclease-free water was added to the pellet, and the tube was warmed in a 55 ºC water bath for 15 minutes.

Section 5 – RNA quantification

RNA quantification was performed using a NanoDrop OneC Microvolume UV-

Vis Spectrophotometer (Thermo Scientific). RNA samples were briefly vortexed to ensure homogeneity, then 1.5 µL of the solution was analyzed with the instrument. The

software within the NanoDrop machine calculated RNA concentrations in µg/µL, by utilizing the Beer-Lambert law:

[RNA] = A260/(ε * b)

Where the molar extinction coefficient (ε) was 0.025 (µg/ml)-1cm-1 and the pathlength

(b) was determined automatically by the software to be between 1.0 mm and 0.03 mm.

The software was also able to identify if DNA or phenol contamination of the RNA sample had occurred during RNA isolation by looking at the ratio of the absorbance at

260 nm (A260) to the A280 or A230. Phenol in the TRI reagent would cause an increase in the absorbance of the samples at 280 nm and 230 nm, so a lower A260/A280 suggests phenol contamination. The software was able to adjust the calculated concentrations based on these contaminants, in order to provide a corrected RNA concentration, which was used in calculations for cDNA creation.

Section 6 – RT-PCR to make cDNA

Reverse transcriptase polymerase chain reaction (RT-PCR) was utilized to create a complementary DNA (cDNA) library from the isolated messenger RNA (mRNA). By using equal masses of total RNA to create cDNA libraries, it becomes possible to compare the level of gene expression in different samples. The mass of RNA used to create the cDNA will sometimes be limited by the RNA sample with the lowest concentration.

To start the RT-PCR, a Moloney Murine Leukemia Virus master mix (M-MLV

MM) was created using M-MLV Reverse Transcriptase (M-MLV RT), M-MLV RT 5X buffer, dNTP mix, and Oligo(dT)15 Primer mix (all from Promega) in a 0.2-mL Thin-

walled Tube with Flat Cap (Thermo Scientific). This M-MLV MM was then combined with nuclease-free water and between 1 and 8 µg of RNA per 50 µL to yield the following final concentrations: 200 units of M-MLV RT, 1X M-MLV RT buffer (50 mM

Tris-HCl, 7 mM MgCl2, 40 mM KCl, 10 mM dithiothreitol), 0.025 mM dNTP mix, 0.025 mM Oligo(dT)15 Primer mix, and between 0.02 and 0.16 µg/µL RNA.

This reaction mixture was subjected to cDNA synthesis using an Eppendorf

Mastercycler pro S with an Eppendorf vapo.protect heated lid. The thermocycler was programmed to start at 25.0 ºC for 10 minutes, 42 ºC for 50 minutes, and then 4 ºC to hold until cancelled. The cDNA samples were stored in a -20 ºC freezer until they were needed for future experiments.

Section 7 – End-point PCR

Polymerase chain reaction (PCR) is a technique that allows targeted amplification of a region of cDNA through the use of specific primer pairs. The concentration of the

PCR product is related to the initial mRNA concentration of the gene of interest, and thus transcription levels of genes can be compared across different cDNA samples. A list of the primers used can be found in Table 1 (Integrated DNA Technologies). End-point

PCR utilizes gel electrophoresis to separate and visualize PCR products after their amplification is complete.

Each end-point PCR reaction mixture was made by mixing 12.5 µL GoTaq Green

MasterMix 2X (Promega), 9.5 µL of nuclease-free water, 1 µL each of 10 µM sense and antisense gene-specific primers, and 1 µL of cDNA. The reaction mixture was then placed into the Eppendorf Mastercycler pro S for amplification of the target sequence.

The thermocycler was set to denature the DNA at 95 ºC for 10 minutes, then cycle through a denaturing step at 95 ºC for 30 seconds, an annealing step at a primer-specific annealing temperature for 45 seconds, and then an elongation step at 72 ºC for 45 seconds that allows for primer extension. These cycles were repeated a set number of times to amplify the gene of interest. Following the cycles, a final 10 minute elongation step at 72

ºC was performed, and then the thermocycler decreased the temperature to 4 ºC and held until cancelled.

Gene (Amplicon Size) Primer Type Tm (ºC) Primer Sequence (5' - 3') LPL Sense 51.7 GGAATGTATGAGAGTTGGGT (308 bp) Antisense 53.6 GGGCTTCTGCATACTCAAAG HKII Set 1 Sense 61.2 AGTACATGGGCATGAAGGGCG (193 bp) Antisense 56.3 ACATCCAGGTCAAACTCCTCTC HKII Set 2 Sense 58.4 GTGGTGAATGACACAGTTGGGA (109 bp) Antisense 57.4 TCTCTTCCATGTAGCAGGCGTT PFKM Sense 55.3 GTGACCAAAGACGTGACCAA (221 bp) Antisense 55.3 AGGCCAATCCTCACAGTAGA β-actin Sense 60.2 TCATGAAGTGTGACGTTGACATCCGT (285 bp) Antisense 60.6 CTTAGAAGCATTTGCGGTGCACGATG Table 1. Primers used for PCR reactions.

Once amplification was complete, the PCR products were loaded onto a 2% agarose gel submerged in 1X TAE buffer (400 mM Tris-acetate, 1 mM EDTA). The gel was made by adding 0.7 mg of agarose (Fisher Scientific) to 35 mL of the 1X TAE buffer, microwaving until fully dissolved, and then adding either 1.75 µL of 10 mg/mL ethidium bromide (Invitrogen) or 3.5 µL of GelRed Nucleic Acid Stain, 10,000X in water

(Biotium) to enable visualization of the PCR products. GelRed and ethidium bromide are small molecules that intercalate into DNA and fluoresce when exposed to UV light.

Images of the gel were captured using a UV transilluminator with a digital camera

(BioRad Quantity One 4.6.6, ChemiDoc XRS, UV Transilluminator Universal Hood or

BioRad ChemiDoc Imaging system with Image Lab Touch software), and intensity analysis was performed using Image Lab software. The wild-type band for each gene was used as a reference to give the relative band intensities for the KD bands in terms of the percentage of wild-type expression. The relative band intensities for the genes of interest were then normalized by dividing by the relative band intensities seen for the β- actin bands.

Section 8 – Quantitative PCR

Quantitative PCR (qPCR) is a technique that utilizes fluorescence and spectrophotometry to determine the concentrations (either absolute if using a standard curve or relative to another sample) of PCR products while amplification is taking place.

A dye that produces fluorescence upon intercalation with double-stranded DNA (dsDNA) is used in the reaction mixture, and fluorescence is constantly monitored throughout the cycles of PCR amplification. The number of cycles required to amplify enough DNA to reach a specific threshold level of fluorescence is called the CT value. This CT value is inversely proportional to the cellular expression of the gene of interest. Therefore, a lower CT value indicates that there was more transcribed mRNA for the gene of interest present during cDNA creation.

Theoretically, the dsDNA created during PCR amplification initially doubles in concentration each cycle. While this is occurring, the amplification is said to be in the

“log phase.” However, due to instrumental limitations, qPCR is unable to detect the difference in the amount of DNA at higher concentrations, so the amplification appears to plateau. The fluorescence threshold is chosen to measure the newly created dsDNA

while it is still in the log phase of amplification, because minute changes in initial cDNA concentrations are detectable.

The qPCR master mix for each reaction consisted of 15 µL of Maxima SYBR

Green/ROX qPCR Master Mix (Thermo Scientific), 10 µL nuclease-free water, and 1 µL each of sense and antisense primers for the gene of interest (10 µM, as in Table 1). Next,

22.5 µL of the qPCR master mix and 2.5 µL of cDNA were mixed to create the reaction mixture, and the reaction mixture was transferred to a Cepheid SmartCycler PCR tube.

The tubes containing the reaction mixtures were placed in a Cepheid SmartCycler II for

PCR amplification with simultaneous fluorescence monitoring. The program used began with a denaturation step at 95 ºC for 120 seconds, then went through 45 cycles of the following: denaturation at 95 ºC for 15 seconds, annealing at 60 ºC for 30 seconds, and elongation at 72 ºC for 30 seconds. After the 45 cycles, a final elongation step at 72 ºC was performed for 120 seconds, followed by a melting curve analysis. The melting curve protocol started the samples at the annealing temperature of 60 ºC, and increased the temperature to 95 ºC in increments of 0.2 ºC per second. The melting curve was used in order to confirm that the PCR amplification was specific to the DNA targeted by the primers. A melting temperature of higher than 80 ºC is expected for dsDNA, and anything lower than that suggests nonspecific amplification or insufficient time for primer extension. The CT values obtained were used to calculate the fold-change between the WT and KD samples through the equation25:

Fold-change = 2-(CT target – CT β-actin)KD – (CT target – CT β-actin)WT

This equation normalizes the CT value of the gene of interest to that of the housekeeping gene β-actin. The fold-change was then multiplied by 100% in order to get the percent expression of the LPL-KD samples relative to the WT.

Section 9 – L6 cell lysis for protein isolation

In order to obtain proteins for analysis, the L6 cells were differentiated as previously described. Following the second day of incubation in differentiation medium

2, the medium was aspirated, and the cells were washed with 5 mL of cold PBS. A stock lysis buffer containing 50 mM Tris-HCl, 2 mM CaCl2, and 0.1% Triton-X100 was created. Working lysis buffer was created by adding 0.01 M concentrations of the protease inhibitors benzamidine, phenylmethanesulfonyl fluoride (PMSF), and aprotinin to the stock lysis buffer immediately prior to each lysis. The flask containing the washed, differentiated cells was then put on ice, and 500 µL of working lysis buffer was added to the cells. The cells were scraped into the lysis buffer in order to facilitate cell lysis and maximize protein yield, and then the resulting cell lysate mixture was transferred to a 1.5- mL microcentrifuge tube. The cell lysate was then centrifuged at 15,000 rpm for 1 minute in order to form a pellet of the insoluble cell materials, and the supernatant was used for protein quantification and experiments. Cell lysate was stored at -20 ºC when not in use, and centrifugation was repeated after each freeze-thaw cycle prior to use in experiments.

Section 10 – Protein quantification assay

Proteins were quantified using a Pierce BCA Protein Assay Kit (Thermo

Scientific). The BCA assay utilizes the ability of proteins to reduce Cu2+ to Cu+ in

alkaline environments, specifically due to reduction interactions with cystine, cysteine, tryptophan, tyrosine, and peptide bonds.26 The extent of the reduction of the copper II ions is related to the concentration of protein in the sample. The Cu+ interacts with bicinchoninic acid (BCA) to produce a BCA-Cu+ complex, which has an intense purple color with a peak absorption at 562 nm. Thus, the BCA assay creates a relationship between the intensity of the absorbance of a sample at 562 nm and the total soluble protein concentration in that sample.

A standard series of bovine serum albumin (BSA) with concentrations of 0.05,

0.10, 0.15, 0.20, and 0.25 mg/mL was created from a stock solution of 2.0 mg/mL BSA in a 0.9% aqueous NaCl solution (Thermo Scientific) by diluting with the same protein lysis buffer or assay buffer used in the cell lysates. The lysate samples were also diluted with the same buffer to between 1:10 and 1:30 of their original concentrations to ensure that they fell within the range of the standard curve. BCA working reagent was prepared by mixing 50 parts Reagent A with 1 part Reagent B, and then 100 µL of the dilute standard, sample, or blank (just lysis buffer) was mixed with 2 mL of BCA working reagent in a glass test tube. The tubes were mixed by vortexing, covered with Parafilm, and then incubated at 60 ºC for 30 minutes. Following incubation, the samples were allowed to cool to room temperature, transferred to a 96-well vinyl assay plate (Corning costar), and measured for absorbance at 562 nm in a BioTek Synergy 2 Multi-Mode

Microplate Reader. The absorbance values of the standards were then used to make a standard curve, from which the unknown protein concentrations were calculated.

Section 11 – Western blot

Western blotting was used to verify that the differences in gene expression between WT and LPL-KD samples that had been observed using PCR also corresponded to different protein levels in the samples. The first step of western blotting involves the separation of proteins via sodium dodecyl sulfate – polyacrylamide gel electrophoresis

(SDS-PAGE). The main protein separation during this technique takes place in the resolving gel, which is typically a higher percentage of acrylamide than the upper, stacking gel. The stacking gel is poured directly on top of the polymerized resolving gel, and is used to ensure that proteins load onto the resolving gel evenly across each of the lanes that the samples will run in.

Resolving gels of 10% (or 12%) polyacrylamide were made by mixing 4.1 mL of ddH2O (3.4 mL for 12% gel), 3.3 mL of a degassed solution of 29.2% acrylamide – 0.8%

N’N’-bis-methylene-acrylamide (bis-acrylamide) (4.0 mL for 12% gel), 2.5 mL of 1.5 M

Tris-HCl buffer at pH 8.8, 100 µL of 10% w/v sodium dodecyl sulfate (SDS), 75 µL of

10% ammonium persulfate (APS), and 7.5 µL of N,N,N’,N’-tetramethylethylenediamine

(TEMED). The resolving gel was poured into a cast with 1.5 mm spacers, and allowed to polymerize for 1 hour.

A 4% polyacrylamide stacking gel was used regardless of resolving gel concentration, and was made by mixing 6.1 mL of ddH2O, 1.3 mL of degassed 29.2% acrylamide – 0.8% bis-acrylamide, and 2.5 mL of 0.5 M Tris-HCl buffer at pH 6.8. This mixture was degassed in a desiccator for 15 minutes, and then 100 µL of 10% APS and

12.5 µL of TEMED were added immediately prior to pouring the gel. A 10-well comb was inserted into the cast, and the stacking gel was allowed to polymerize for 90 minutes.

The SDS-PAGE method is useful for separating denatured proteins by protein size. Protein samples are prepared in Laemmli reducing buffer, which utilizes SDS and

2-mercaptoethanol to denature proteins and reduce disulfide bonds in protein samples.

Samples were prepared in 0.2-mL microcentrifuge tubes by adding 10 µL of 4X Laemmli reducing buffer to volumes of WT or LPL-KD cell lysate that yielded an equal mass of protein. The total sample volumes were then brought up to 40 µL with ddH2O. The tubes containing the samples were then submerged in boiling water for 5 minutes, and allowed to cool to room temperature. Next, 35 µL of the samples or 5 µL of EZ-RUN Pre Stained

Protein Marker (Fisher Scientific) were loaded into the wells formed in the stacking gel.

Electrophoresis was carried out at 200 V in a Mini-PROTEAN Tetra System (Bio-Rad) filled with a sufficient amount of electrode buffer (25 mM Tris, 192 mM glycine, 0.1%

SDS) and continued until the dye front ran off of the gel (approximately 1 hour).

Following electrophoresis, the gel was removed from the electrode buffer and separated from the glass plates. The stacking gel was cut off with a razor blade and discarded. The resolving gel portion, which contained the proteins of interest, was then set up to transfer to a polyvinylidene fluoride membrane (Immobilon) using the Mini

Trans-Blot Electrophoretic Transfer Cell (Bio-Rad). The gel and membrane were set up in a transfer cassette and submerged in transfer buffer (25 mM Tris, 192 mM glycine,

15% methanol, at pH 8.6), then run either overnight at 30 V, or for 1 hour at 100 V. The membrane, which at this stage contained the adsorbed proteins, was then cut with a razor blade to be closer to the original size of the gel, and incubated with rocking at 37 ºC in blocking buffer (20 mM Tris, 150 mM NaCl, 2 mM CaCl2, 1% gelatin, 0.05% TWEEN-

20, adjusted to pH 7.4) for between 2 and 16 hours. This blocking step was performed in

order to block nonspecific binding of antibody to the membrane. The membrane was then removed from the blocking buffer, and washed twice for 5 minutes each with 1X wash buffer (20 mM Tris, 150 mM NaCl, 2 mM CaCl2, 0.05% TWEEN-20).

Next, the membrane was incubated for 2 hours in primary antibody diluted with blocking buffer with constant rocking. A 0.4 µg/mL solution of HXKII (B-8) mouse monoclonal IgG2a (sc-374091, Santa Cruz Biotechnology) in blocking buffer was used as the hexokinase II primary antibody solution. Following incubation with the primary antibody, the membrane was washed with wash buffer 4 times for 5 minutes each, and then incubated in the secondary antibody solution for 1 hour. The secondary antibody solution used for hexokinase II was a 32 ng/mL solution of ImmunoPure Antibody Goat

Anti-mouse IgG (H+L) with a horseradish peroxidase (HRP) label (cat # 31430,

Pierce/ThermoFisher) diluted in blocking buffer. Table 2 contains information about the primary and secondary antibodies used for each protein of interest.

Protein Primary Antibody Secondary Antibody 0.4 µg/mL HXKII (B-8) Mouse 32 ng/mL ImmunoPure Goat Anti- Hexokinase II Monoclonal IgG2a * mouse IgG (H+L), HRP label ** 1 µg/mL ms mAb to beta Actin, 1 µg/mL Rb pAb to ms IgG (HRP), β-actin ab8226 *** ab6728 *** 0.4 µg/mL PFK-1 (G-11) Mouse 32 ng/mL ImmunoPure Goat Anti- PFK-1 Monoclonal IgG2b * mouse IgG (H+L), HRP label ** Table 2. Antibodies used in western blotting. *=Santa Cruz Biotechnology, **=Pierce/ThermoFisher, ***=abcam.

The membrane was washed with wash buffer 5 times for 5 minutes each following the secondary antibody incubation. It was then incubated in the dark for 5 minutes in SuperSignal West Pico PLUS Chemiluminescent Substrate (ThermoFisher

Scientific), which is a sensitive, luminol-based chemiluminescent substrate used to

facilitate the detection of the HRP that was conjugated to the secondary antibodies. For the β-actin western, the reagents used were Detection Reagent 1, Peroxide Solution (prod

# 1859700, Thermo Scientific) and Detection Reagent 2, Luminol Enhancer Solution

(prod # 1859697), which were mixed and incubated with the membrane for 2 minutes in the dark. Visualization and imaging of the membrane was then carried out using a Bio-

Rad ChemiDoc Imaging System with Image Lab Touch Software on the setting

“Chemiluminescent Blot 647SP, No Light”. The images were processed and quantified using the programs GIMP 2, Image Lab software, and Image J software. The intensities of the bands in the hexokinase western blot were normalized with the intensities of the corresponding β-actin bands for each sample. The intensity of the HK band was divided by that of the β-actin band for this normalization.

Section 12 – Hexokinase activity assay

A Hexokinase Activity Assay Kit (Colorimetric) from abcam was used to measure and compare the activity of the hexokinase enzyme in WT versus LPL-KD cells.

Hexokinase converts glucose into glucose 6-phosphate (G6-P), which can then be oxidized by glucose 6-phosphate dehydrogenase (G6PD) into 6-phospho-D-glucono-1,5- lactone (6PG). The oxidation of G6-P is carried out alongside the reduction of the coenzyme NAD+ to produce NADH (Figure 8). This HK assay kit utilizes a colorless probe that becomes reduced by NADH to yield a colored product with a strong absorbance at 450 nm. Thus, spectrophotometry that measures absorbance at 450 nm allows for determination of NADH concentration. A comparison of the absorbance values at different time points can be used to measure the rate of formation of NADH, and in turn provides insight into the hexokinase activity in each sample.

The samples used for the HK activity assay were prepared by lysing cells in the

HK assay buffer provided with the kit. In order to do this, cells were differentiated the same way as previously described, then washed twice with cold PBS. Next, 8 µL of HK assay buffer per cm2 of adherent surface area was added to the cells, and the cells were scraped into the buffer to yield HK assay lysate. The HK assay lysate was transferred to a 0.6-mL microcentrifuge tube and centrifuged at 12,000 rpm for 5 minutes. The supernatant was collected and transferred to a clean microcentrifuge tube, which was stored at -20 ºC until ready to use in the assay.

Figure 8. Theory behind the hexokinase activity assay. D-glucose is the original substrate converted to G6-P by hexokinase. NADH produced in the subsequent reaction reacts with the probe to yield a colored product. The protein concentration in the HK assay lysates was determined using the BCA assay as described previously, except HK assay buffer was used in place of protein lysis buffer.

Standards were created for the HK activity assay by reconstituting a stock solution of 1.25 mM NADH Standard and diluting it to final concentrations of 0, 2.5, 5.0,

7.5, 10.0, and 12.5 nmol/well. The reaction mixtures were set up in a 96-well vinyl assay plate to be a 1:1 ratio of the diluted sample/standard/positive control (HK-II from

Bacillus sp.) to reaction mix or background reaction mix. The reaction mix consisted of

HK assay buffer, enzyme mix (which most likely contains G6PD), developer, coenzyme

(probably NAD+), and hexokinase substrate (D-glucose). The background reaction mix was the same as the reaction mix, except no hexokinase substrate was added, and more

HK assay buffer was used instead. The exact concentrations and contents of each component are proprietary and unknown.

Once the samples and standards had been mixed with either the reaction mix or background reaction mix, they were allowed to incubate at room temperature in the dark for 20 minutes, and then the absorbance at 450 nm was read in 5 minute intervals for the next 40 minutes. The blank absorbance (that of the 0 nmol/well standard) was subtracted from all of those raw sample values and the standards to yield the corrected absorbance for each well. The corrected absorbance values of the standards were then used to construct a standard curve, which could be used to determine the concentration of NADH in the samples. These NADH concentrations were then divided by the mass of total protein (in mg) added to each sample in order to obtain the nmol NADH per mg of protein. This NADH/mg protein data was plotted versus time to yield information about the rate of NADH formation in each sample. The linear range was determined for each sample, and a best-fit line was determined for the data. The slope of the linear fit that was applied to the graph gave nmol of NADH produced per mg of protein added per minute, and the y-intercept corresponded to initial nmol of NADH in the lysate at the start of the assay. In order to more directly compare the rate of NADH formation, we wanted to remove this initial NADH from our calculations. To this end, the y-intercept

calculated for each sample was subtracted out from each of the nmol NADH/mg protein values for that sample to yield background-corrected NADH production values.

NADH baseline assay

A modified version of this procedure was performed in order to measure the rate of NADH formation in the samples in the absence of any additional coenzymes or enzymes supplemented by the assay kit. For this NADH baseline assay, 50 µL of KD sample was added to 48 µL of assay buffer with 2 µL of developer. The developer would thus probe for the NADH that was present in the cells, and any change in the absorbance at 450 nm over time could be attributed to NADH formation from enzymatic reactions taking place in the lysate. The absorbance was read initially, then every 10 minutes for

30 minutes. The absorbance of the samples increased at a dramatically higher rate than that of the standards, which justifies the subtraction of the y-intercept as the background in the main assay data analysis.

Concentration of assay lysates

In an effort to reduce the background baseline NADH production, HK assay lysates were concentrated using a Centricon-3 microconcentrator (Amicon). The microconcentrator works by ultrafiltering the lysate sample through an anisotropic membrane with a molecular weight cutoff of 3 kDa. Solvents and solutes with a lower molecular weight than the cutoff pass through the membrane and are discarded, leaving behind concentrated enzymes with far less endogenous glucose, NAD, and NADH.

Due to the increased volume of lysate required for microconcentration, the differentiated cells were lysed in 16 µL of HK assay buffer per cm2 of flask, but

otherwise collected under the same conditions as described previously. The microconcetrators were assembled in concentration mode according to the manufacturers operating instructions (Publication I-259, revision J), and 1 mL of lysate was added to each microconcentrator. The assembly was centrifuged at 4,000 rcf for 90 minutes at 4

ºC in centrifuge with a fixed-angle rotor (Beckman Coulter Avanti J-E Centrifuge), which led to a reduction of sample volume to about one-third of the original volume. Next, 500

µL of HK assay buffer was added to each sample, and the samples were centrifuged for an additional 50 minutes under the same conditions. Finally, the filtrate was discarded, the sample reservoir was inverted, and the samples were centrifuged in this recovery mode at 500 rcf for 2 minutes to transfer all of the concentrated retentate into the retentate cup. The retentate was then used in place of the lysate samples during the hexokinase activity assay procedure described previously.

The data from the concentrated hexokinase assay was processed in a manner that was closer to the recommended data analysis method suggested in the HK assay manual.

A standard curve was made using the standards with known NADH concentrations, and used to determine the [NADH] in each sample or background sample. As previously described, the samples were prepared with reaction mix, and the background samples

(BG samples) were prepared using background reaction mix. The nmol of NADH calculated this way for each sample or BG were then divided by the mg of total protein added to each reaction well from the concentrated lysates, in order to yield the nmol of

NADH per mg of protein. Then, the calculated nmol NADH/mg protein for each BG sample was subtracted out from the corresponding sample in order to determine the background-subtracted NADH present in the sample, again in units of nmol NADH/mg

protein. These background-subtracted NADH values were then plotted versus time, and the slope of the linear regression provided information about the HK activity by looking at the nmol of NADH produced per mg of protein added per minute (nmol/(mg*min)).

Section 13 – Statistical analysis

Statistical significance of data was determined using the Student’s t-Test, as calculated by the function “TTEST” in Microsoft Excel. The method used was a one- tailed, paired data t-Test. When the data points were not treated as paired, but rather as two-samples with equal variance, the calculated P value decreased further, thus the data would be even more significant than reported if analyzed in that manner. P values less than 0.05 were considered significant. The standard deviations and averages of the data were also calculated using Microsoft Excel. These standard deviation values are what are reported as the uncertainty when discussing the averages of multiple trials.

Chapter 3 – Results

Section 1 – Morphology of LPL-KD cells

The wild-type L6 cells and the LPL-knock-down L6 cells exhibited slightly different morphology and growth rates. Figure 9 shows images taken of the cells both before and after differentiation. The LPL-KD cells divided more rapidly in the growth medium, and appear to be less elongated and less aligned with one another following differentiation. They also continued to divide for longer in the differentiation medium than the WT L6 cells.

Figure 9. Comparison of the morphology of the L6 WT vs LPL-KD cells. a) Growth-phase WT cells. b) Growth-phase LPL-KD cells. c) Differentiated WT cells. d) Differentiated LPL-KD cells.

Section 2 – Effects of LPL-KD on RNA expression levels

PCR array data that had been collected by a previous student in our lab

(unpublished) suggested that there was a difference in the expression of several genes involved in glycolysis as a result of LPL knock-down. The expression of the genes that encode hexokinase II (HKII gene) and phosphofructokinase-1 (PFKM gene) were of particular interest, as these enzymes are required for irreversible, energy dependent steps in the glycolytic pathway. Regulation of these genes would thus have a significant impact on the glucose metabolism of a cell. Gene expression was measured through the use of end-point PCR, as shown in Figure 10.

Figure 10. End-point PCR products separated via 2% agarose gel electrophoresis. PCR amplification was carried out for 30 cycles with an annealing temperature of 60 ºC. β-actin was used as a housekeeping gene to show equal cDNA loading. *Relative intensities were normalized such that β-actin was equal to 100%. The end-point PCR confirmed LPL knock-down, as there was no detectable LPL band for the LPL-KD cells. This LPL knock-down increased the expression of the HKII gene (which encodes for a monomer of hexokinase II) to 169% of WT levels, while

decreasing the expression of the PFKM gene (which encodes for a monomer of phosphofructokinase-1) to 93% of WT levels. The end-point PCR also revealed that there was no nonspecific amplification of cDNA by the primers that were used, as there was only one observed band in each lane. In order to further investigate gene expression, qPCR was performed using the same primers (Figure 11).

Regulation of Gene Transcription in LPL-KD Cells

300 253* 250

200

150

100 72.1* 50 Percent (Relative to WT)to (Relative Percent 0.02** 0 LPL HKII PFKM Gene of Interest

Figure 11. Regulation of gene transcription in LPL-KD cells as determined using qPCR. The experiment was performed in triplicate (n=3). *P<0.01, **P<1x10-7. The data obtained from the quantitative PCR supports the trends that were observed in the end-point PCR. In L6 cells that had LPL expression knocked down to trace levels, hexokinase II expression was upregulated to 253±22% of wild-type levels.

In the same cDNA, the expression of the PFKM gene was downregulated to 72.1±5.3% of the WT expression levels. The difference in the gene expression is more pronounced in the qPCR data than in the end-point data due to the higher sensitivity of the technique.

Section 3 – Effects of LPL-KD on protein expression levels

In order to further explore the impact of knocking down LPL in rat skeletal muscle cells, we shifted focus from gene transcription towards protein expression. In theory, the mRNA that was used to make the cDNA library would normally act as a template for protein translation, so it was expected that hexokinase II protein levels would be higher in the LPL-KD cells than in the WT cells. SDS-PAGE followed by western blot analysis was used to test this hypothesis (Figure 12).

The bands corresponding to the LPL-KD lysate samples were wider and more intense than those of the WT cells due to the increased concentration of HK II protein contained in the KD sample lysates (Figure 12a). In order to ensure that this pattern was not due to a difference in the amount of total protein loaded, the bands were normalized to a β-actin western blot of the same proteins (Figure 12b). These normalized values can be found in the table, labeled as Figure 12c. The normalized density of the KD 2 band was 278% higher than that of WT 3, while the WT 1 band was 54% of the normalized intensity. The normalized intensity of the KD 4 band, at 4.94, is the furthest from all of the other intensities, and is almost four times as high as that of the KD 2 band. This value greatly impacted the calculated average, but there is not enough data to determine which, if either, of the KD results was statistically insignificant. The average relative intensity of the corrected WT protein lysates was 0.36 ± 0.15, and the average of the

LPL-KD samples was 3.11 ± 2.59. All of the lanes had a thin, faint band at a slightly higher molecular weight than the main hexokinase II band. This could be due to post- translational modification of the enzyme, or potentially primary antibody binding to a different hexokinase isoform that is slightly larger in mass.

12 c)

Sample Normalized Intensity Sample Normalized Intensity WT 1 0.25 KD 2 1.28 WT 3 0.46 KD 4 4.94

Avg. 0.36 Avg. 3.11 Std. Dev. 0.15 Std. Dev. 2.59

Figure 12. Western blot analysis of HK II protein concentration in L6 cells. a) Western blot image using anti-HK II antibodies. b) Western blot using anti-β-actin antibodies to be used as a protein loading control. c) Table of HK II intensities normalized against the β-actin intensities, including the averages for each cell type.

Section 4 – Effects of LPL-KD on hexokinase activity

Fatty acids, particularly long-chain acyl CoA (LCACoA), have been shown to have regulatory effects on glycolysis through inhibition of the enzymes hexokinase,

glucose 6-phosphatase, and pyruvate dehydrogenase.20 Lipoprotein lipase facilitates the hydrolysis of triglycerides into glycerol and free fatty acids, so it would follow that

LCACoA levels would be higher in wild-type L6 cells than in LPL-KD L6 cells. The experiments conducted to determine the relative expression of HK II have indicated that there is an increased expression of the enzyme in the LPL-KD cells, but we were also interested to see if there was a difference in the enzyme activity between the WT and

LPL-KD cell lines.

A hexokinase activity assay was performed on cell lysate from 5 flasks of differentiated WT and LPL-KD L6 cells. The activity assay utilized the absorbance of a probe+NADH complex at 450 nm to determine the activity of hexokinase in the samples.

In order to interpret the absorbance values as NADH concentrations, a standard curve was made using the reaction mixture and NADH standards of known concentration

(Figure 13).

HK Assay Standard Curve 0.8 0.7 0.6 0.5 0.4 y = 0.0563x - 0.0165 0.3 R² = 0.9881 0.2

Absorbance nm) (450 0.1 0 0 2 4 6 8 10 12 14 -0.1 NADH (nmol)

Figure 13. Standard curve used for the hexokinase assay data analysis.

The sample NADH concentrations obtained using the standard curve needed to be corrected for the mass of protein used in each assay well. The L6 cells are adherent cells, and the LPL-KD L6 cells tend to grow at a higher rate than the WT cells. For this reason, the different flasks of cells grown often had different total amounts of cells following the

5 day incubation required for differentiation, so it was not possible to confidently analyze activity “per million cells.” The sample lysates thus had different initial concentrations of protein, so it was important to normalize the NADH production to the level of total protein in the lysate. This necessitated the use of protein concentration assays, which were used to determine the mass of total protein added during the activity assay.

We suspected that the activity of the hexokinase in the WT cells would be lower than that of the LPL-KD cells, due to the increased potential for inhibition of the enzyme by LCACoA. However, our data suggests a decrease in HK activity in the KD samples

(0.57±0.18 nmol NADH/(mg*min)) compared to the WT samples (0.75±0.025 nmol

NADH/(mg*min)) (Figure 14).

Figure 14. Hexokinase activity in WT and LPL-KD L6 cells. The values shown are in units of nmol of NADH per mg of protein per minute. WT flasks were numbered with odd numbers, while KD flasks were even numbers. The average was determined with n=5 and P<0.05.

The apparent rates of NADH formation were calculated as the slope of the line when nmol of NADH per mg of protein was plotted versus time for each individual sample. The slopes seemed fairly consistent in WT samples but variable in the KD samples. However, the background (assay performed in the absence of added glucose, the substrate for HK) absorbance values were dramatically different between various samples. In order to correct for this background difference and allow for more direct comparison between WT and LPL-KD samples, the y-intercept of the linear fit of each sample was used as the background (NADH per mg of protein at time zero) and subtracted from each of the nmol NADH/mg protein values for that sample. These y- intercept-subtracted values are shown together in Figure 15.

Intercept-subtracted NADH/mg vs Time for all Samples 35 30 25 20 15

10 NADH formed NADH (nmol/mg protein) (nmol/mg 5 0 0 5 10 15 20 25 30 35 40 45 Time (min)

WT 1 KD 2 WT 3 KD 4 WT 5 KD 6 WT 7 KD 8 WT 9 KD 10

Figure 15. Comparison of the nmol of NADH formed per mg of protein per minute for each sample. The WT samples were depicted as circles in shades of blue, and the KD samples were depicted as squares in shades of orange.

To further investigate the variation in initial NADH concentration between samples, a NADH baseline assay was performed on the LPL-KD samples to estimate the

level of endogenous NADH production (Table 3). In this assay, only the developer

(which contained the colorimetric probe) was added to the samples, without adding any additional enzyme substrate, coenzyme, or supplementary enzymes to the lysates. This assay was designed to test for all NADH that was originally present in the lysate or was formed due to substrates and dehydrogenases already present in the samples (cell lysates). The assay did not specifically target hexokinase-dependent production of

NADH.

NADH KD Baseline nmol/(mg*min) 2 0.04 4 0.43 6 0.87 8 0.94 10 0.82 Table 3. Baseline NADH formation in HK assay samples. The HK assay samples produce NADH in the absence of supplemental assay reagents, as determined by a change in A450. This absorbance change was due to one or more components of the cell lysate as it was not observed for the NADH standards. Absorbance values were measured for the first 30 minutes of the reaction. This baseline level of NADH formation varied greatly between the different LPL-

KD samples, with a maximum of 0.94 nmol/(mg*min) for KD 8 and a minimum of 0.04 nmol/(mg*min) for KD 2. These same samples correspond to the maximum and minimum NADH values observed during the activity assay, so the baseline NADH formation is likely a significant source of error for the HK assay.

These baseline values are helpful to the understanding of issues that have complicated the analysis of HK activity, but cannot be used directly to correct the HK activity assay data. The baseline rates of NADH formation correspond to absorbance values measured during the first 30 minutes of the interaction between the samples and the developer probe. However, the actual HK assay started after 20 minutes of

incubation in the dark, so there is limited data overlap. It is possible that the initial rates of NADH formation in the sample would be higher than the rates following 20-30 minutes of incubation.

In an effort to eliminate the high baseline NADH production, three sets of cell lysates were concentrated using a Centricon-3 microconcentrator (Amicon). This method increased the concentration of enzymes larger than 3 kDa, while simultaneously reducing the concentration of endogenous glucose, NADH, NAD, and other possible small molecule contaminants from the lysates. The concentrated retentate samples were used in the hexokinase activity assay, which yielded the absorbance values in Table 4 at 450 nm.

Wild-Type LPL-KD Time (min) Sample BG ΔA450 Sample BG ΔA450 20 0.244 0.233 0.011 0.249 0.249 0 25 0.268 0.264 0.004 0.271 0.269 0.002 30 0.289 0.28 0.009 0.29 0.285 0.005 35 0.31 0.293 0.017 0.304 0.297 0.007 40 0.326 0.304 0.022 0.317 0.307 0.01 45 0.341 0.314 0.027 0.33 0.316 0.014 50 0.354 0.318 0.036 0.341 0.322 0.019 55 0.366 0.323 0.043 0.347 0.327 0.02 60 0.372 0.328 0.044 0.353 0.331 0.022 Table 4. Raw absorbance value comparison of concentrated lysates. The absorbance was measured at 450 nm. BG lysates were treated with the background reaction mix, Sample lysates were treated with reaction mix containing the HK substrate. ΔA450 is the difference between Sample and BG absorbance. The method of data analysis that was suggested in the instructions for the HK assay involves subtracting the background absorbance out from each sample absorbance to get the ΔA450, then subtracting the absorbance of the blank standard from that difference. However, the blank standard in this case had an absorbance of 0.077, which

is larger than any of the ΔA450 values obtained for this assay. This meant that the difference between the absorbance values was too small to be fit to the standard curve for analysis of NADH concentration.

Instead, the absorbance of the blank was subtracted out from all sample and background absorbance values, and the nmol of NADH in each was determined using the standard curve. Then, the samples were corrected for protein concentration and subtracted by the corrected BG samples. These background-subtracted nmol of NADH per mg protein values were then plotted versus time in order to find the rates of NADH production that were solely dependent upon hexokinase activity (Figure 16).

NADH Formation in Concentrated Samples 6.00 y = 0.2889x - 6.3597 5.00 R² = 0.9907 4.00

3.00

2.00 NADH NADH formed

(nmol/mg (nmol/mg protein) y = 0.1098x - 2.2392 1.00 R² = 0.9952 0.00 15 20 25 30 35 40 45 Time (min)

LPL-KD WT Linear (LPL-KD) Linear (WT)

Figure 16. Comparison of NADH formation in the concentrated WT and KD samples after background-subtraction. This method of analysis provided NADH formation values of 0.29 nmol/(mg*min) for the concentrated WT sample, and 0.11 nmol/(mg*min) for the concentrated KD sample. Based on these results, it appears that hexokinase activity is in fact higher in the wild-type lysate than in the KD lysate, which is the opposite of what

was seen as the trend in gene and protein expression. The implications of this interesting observation are further discussed in the next section.

The observed impact of knocking down LPL in rat skeletal muscle cells is summarized in Figure 17. All of the claims regarding increasing or decreasing are relative to the WT L6 cell line.

Figure 17. Summary of the main findings of this research. The hexokinase II regulation has been studied fairly extensively, but there is more to explore about PFK-1.

Chapter 4 – Discussion

The purpose of this project was to explore the effects of reducing LPL expression in muscle cells on the glycolytic enzymes hexokinase II and phosphofructokinase-1. We were interested in this subject because previous studies had shown that glucose utilization was increased in LPL-knock-down cells compared to their wild-type counterparts. To further investigate this observation, we compared: (i) the gene transcription levels for

HKII and PFKM, (ii) the relative quantity of hexokinase II protein, and (iii) the catalytic activity of hexokinase II enzyme in the WT and LPL-KD L6 cell lines.

Section 1 – Effect of LPL on gene expression

Our data showed that the expression of the HKII gene and the PFKM gene were altered in LPL-KD L6 muscle cells. Interestingly, the absence (or substantial reduction) of LPL production in the cells appears to have opposite effects on the expression of the two genes. The HKII expression was increased in the LPL-KD cells, while that of PFKM was decreased compared to their WT counterparts (Figures 9 and 10). These trends were observed in both end-point PCR and quantitative PCR, and were consistent across multiple replicate experiments.

There are several reasons why HK II and PFK-1 may be inversely regulated by the knock-down of LPL. One primary difference in the LPL-KD cells compared to the

WT cells is a significant decrease in the availability of free fatty acids for metabolism.

The decrease in LPL products, such as FFAs and monoglycerides, would cause the LPL-

KD cells to be more dependent on glucose metabolism for their energy requirements.

Therefore, it would be beneficial for the cell to upregulate hexokinase expression in order

to increase the amount of glucose that can be partitioned within the cell as glucose 6- phosphate. The conversion of glucose to G6-P is additionally useful because it maintains the concentration gradient of lower intracellular glucose than extracellular glucose. This concentration gradient allows glucose transporters that use facilitated diffusion (i.e.

GLUT4) to continue to function.27

However, an increased demand for glucose uptake in the LPL-KD cells does not necessarily mean that there is a need for an upregulation of PFK-1. Recall that the G6-P that results from the phosphorylation of glucose by HK II can have multiple fates, including use in glycogen synthesis or the pentose phosphate pathway (PPP).4 On the other hand, the product of PFK-1 catalysis, fructose 1,6-bisphosphate, can only be used in glycolysis. Therefore, an upregulation of both HK II and PFK-1 in the cell could potentially lead to a substantial excess of pyruvate and ATP production, which would deplete valuable cellular resources.

The upregulation of the HKII gene and the downregulation of the PFKM gene should have several noteworthy effects on the intracellular concentration of glucose metabolites/glycolytic intermediates. The simultaneous increase in HK II and decrease in

PFK-1 could lead to a buildup of G6-P, which would be alleviated partially via incorporation into glycogen or utilization in the PPP.28,29 However, since G6-P is an allosteric inhibitor of the HK II enzyme, would there be a substantial change in the activity of the enzyme between the different cell lines? We wanted to explore that subject further, but first we thought it was important to analyze the concentrations of the

Hexokinase II protein in our cellular lysates in order to make sure that the enzyme was in fact upregulated in the LPL-KD cells.

Section 2 – Effect of LPL on hexokinase II protein levels

Our comparison of the hexokinase II protein levels using western blotting (Figure

12) seemed to follow a similar trend as the one observed for the variation of gene expression. Following normalization of the HK band intensities to β-actin intensities as a loading control, it was observed that there is an increased concentration of HK II in the

LPL-KD cells when compared to the WT cells. One noteworthy comparison is that of

KD 2 to WT 3. The normalized protein level of HK II in KD 2 was approximately 278% higher than that of WT 3. This is a very close match to the qPCR data that was obtained for the gene expression, which suggested that the HKII gene in LPL-KD cells was upregulated to 253 ± 22% of the wild-type levels (Figure 11).

The other two samples, WT 1 and KD 4, were not as close to the qPCR data, but still supported the trend that was observed by looking at gene expression. The WT 1 hexokinase II band was the faintest, but its β-actin band was similar in intensity to the

KD 2 and WT 3 bands. This led to the WT 1 normalized intensity to be 0.25, which is roughly half of the normalized intensity of WT 3. However, this still supports the finding that there is less HK II in the WT samples. For the lysates from KD 4, the HK western showed the largest band of all, indicating that there was more HK II present in that sample than any of the other three samples. However, our β-actin control lane for KD 4 showed the faintest band, which acted to amplify the intensity difference initially observed in the HK western when it was used in normalization. The combination of the highest HK intensity and the lowest β-actin intensity led to the normalized intensity of

4.94 for KD 4, which is more than tenfold greater than the highest WT intensity.

The averages of the WT and KD sample intensities are reported, but may not be quantitatively reliable due to the small sample size (n=2). The normalized intensity average for the wild-type samples was 0.36 ± 0.15, and the average for the KD samples was 3.11 ± 2.59. It is noteworthy that the uncertainty in the WT is nearly half of the WT average, while the uncertainty from the KD samples was over 80% of the KD average. It is likely that this would be alleviated by having more data sets, which would enable us to determine if any of the normalized values were statistical outliers. However, complications in the western blotting process led to many failed replication attempts, so we were unable to get any data from additional samples prior to the time of submission.

In all of the lanes of the hexokinase II western blot that contained the protein samples, there is a very faint band at a slightly higher molecular weight than the major observed band. The source of this band, which will be referred to as a halo band, is not entirely clear to us at this point. There are several possible explanations that could lead to a slightly higher molecular weight band being observed. One possibility is that the antibody is binding to a different isozyme of hexokinase that is present in the sample.

Three different hexokinase isozymes, hexokinase I-III, each have molecular masses of approximately 100 kDa, with slight variations.17 The halo bands could be a result of the primary antibody binding to a region on one of these isozymes that is similar to the target region of HK II. The most likely isozyme would be hexokinase I, as it is ubiquitously expressed in adult tissues, and is sometimes even utilized as a housekeeping protein for normalization.30

Another possibility for the cause of the halo bands could be post-translational modification of the HK II enzyme. Post-translational modifications, which have been

previously observed in hexokinase, could lead to a higher molecular weight protein from the same sized mRNA.31 A study of hexokinase activity in tumor cells showed that the post-translational modification of the enzyme actually increased its catalytic activity.32 It is unclear whether or not this would be the case for this particular modification, as the band is observed at a similar intensity in both wild-type and LPL-knock-down cells.

Unfortunately, complications with the availability of a reliable antibody prevented research on the protein levels of the phosphofructokinase-1 enzyme in the samples. The western blot attempts carried out using an anti-PFK-1 antibody available in the lab (Santa

Cruz Biotechnology) did not yield any bands, and after multiple trouble-shooting trials, we were unable to determine what parameters needed to be adjusted. The secondary antibody that was used for the successful HK II western blot was theoretically suitable for binding to the PFK-1 primary antibody as well. However, we were never able to observe any distinguishable bands using those secondary antibodies, and additional antibodies that we tried also failed to produce results.

Section 3 – Effect of LPL on the activity of hexokinase

It was clear to us that hexokinase II transcription and translation is increased in

LPL-KD cells, but next we wanted to investigate about the specific activity of the enzyme in the different cell lines. In order to do so, a hexokinase activity assay was performed. The assay utilized a probe that produced a strongly-colored complex in the presence of the cofactor NADH, which was expected to be produced as a result of the oxidation of G6-P by G6PD, a reaction of the PPP (as summarized in Figure 8).

Our initial hypothesis was that the increased availability of free fatty acids in the

WT cells would lead to inhibition of the HK activity, thus lowering the specific activity in WT samples. However, we observed the opposite in the first HK activity assay, as the

LPL-KD cells appeared to exhibit lower HK activity than the WT cells (Figure 14). The standard deviation of the WT samples from that HK assay trial were much smaller than in the LPL-KD samples. In fact, the average for the WT flasks, 0.75 ± 0.025, is actually within one standard deviation of the LPL-KD average of 0.57 ± 0.18. Despite the higher variation among the LPL-KD samples, all but one of the KD activities was lower than any of the WT activities (Figure 15). For this reason, we were fairly confident that the decreased NADH production in the KD samples was not an artifact, but we did run into several experimental issues that we needed to address in order to further support these findings.

The experimental procedure involved the inclusion of separate background (BG) reactions that lacked only one component present in the test samples, the exogenously added substrate (D-glucose). The instruction manual called for the absorbance of background reactions to be subtracted from the absorbance of the test samples. However, the background reaction absorbance values were either nearly equal to or greater than the sample absorbance values, so we were not able to carry out analysis using the background subtraction for the first set of HK assay samples. This issue most likely was caused by excessive levels of glucose in the muscle cell lysate, which may have already been at saturating or near-saturating concentrations for the HK enzymes. As such, the addition of more D-glucose substrate would not substantially raise the activity/rate of catalysis of the HK, so the sample activity would be approximately the same as that of its

background. This issue does not take any other sources of NADH formation into consideration, but it does assume that any other non-HK-dependent NADH formation would be present in both the BG and the sample.

In an effort to see just how much endogenous NADH is already present in or produced by the lysates independently of added substrate, we carried out a NADH baseline assay (Table 3). NADH is the measured product for the reaction, so any extra production or presence of NADH needs to be accounted for. If background subtraction worked, this would theoretically take care of the issue. However, it did not work for us with the initial HK assay, so we wanted to determine just how much of an impact endogenous production had on our data. The baseline assay results clearly indicate that there is a significant contribution of NADH from other sources that is affecting the perceived HK activity, and that background subtraction is critically important to obtain more meaningful data from this assay.

We were not able to directly subtract out data that was obtained in the baseline assay from the HK activity assay data for several reasons. One primary concern was a lack of sufficient quantities of WT sample to perform another assay on. Following protein concentration assays and the HK activity assays performed with the samples, there was no longer enough of the wild-type HK assay lysates remaining to use in the baseline assay. The next issue lies with the experimental differences between the HK assay and the baseline assay. For the HK assay, it was most important to measure the formation of NADH as a result of HK activity, so the initial concentration of NADH and initial rate of NADH production from other sources was not very important. For this reason, the HK assay directions suggested incubation in the dark for the first 20 minutes

after mixing the sample with the reaction mixes, and then measuring the absorbance of the samples at 450 nm at different time points over the following 40 minutes. On the other hand, when we were trying to gain information about the endogenous NADH production, we did not feel a need to incubate in the dark prior to measuring the absorbance. We were interested to see the initial absorbance, as it would likely correspond closely to the initial NADH concentration. After that, we could observe how the absorbance, and thus the concentration, increased over the course of the experiment.

In order to gain data about the initial concentration/rate, we measured the absorbance immediately following the mixing of the sample and the developer solution, and then checked again every 10 minutes for the next 30 minutes. Because of this design, there was limited data overlap between the baseline assay and the standard HK assay, so there was no way to use the baseline assay for normalization or background-subtraction purposes.

The next step in our experimental design was to try and reduce the baseline

NADH production in the samples by concentrating the lysates. This concentration procedure was designed to reduce the concentrations of small molecules and proteins smaller than 3 kDa, which would theoretically reduce the concentration of glucose and substrates responsible for endogenous NADH production, as well as endogenous

NADH/NAD+ already present. The primary goal of this was to reduce the glucose concentration to below the level that saturates the HK enzymes, thus enabling us to have a measurable change in activity between the HK reaction samples and their background samples. As an added benefit, it was possible that the endogenous NADH production by unrelated dehydrogenases would also be reduced due to removal of their substrate

(NAD+) following concentration of the lysates. The raw absorbance data that came from the HK assay of the concentrated samples is shown in Table 4.

Although this concentrated HK assay did seem to improve the difference between the background absorbance and the corresponding sample’s absorbance, it was still not enough to completely follow the manufacturer recommendations for data analysis. It was recommended to just use the difference in the absorbance of the sample and the BG at

450 nm, the ΔA450, to plot on the standard curve and determine the concentration of

NADH in the sample at that time. However, in order to do the analysis that way, the

ΔA450 would need to be larger than the absorbance of the blank (the 0 nmol NADH standard), which was not the case for either of our samples. In an attempt to meaningfully interpret our data, we used the modified analysis method as described in the

Materials and Methods section which allowed us to determine the background-subtracted

NADH values. These values were then used to calculate the rates of NADH production that were the result of glucose-dependent HK activity (Figure 16). Two additional samples for both LPL-KD and WT were concentrated and used in the activity assay, but they did not yield sample absorbance values that were higher than the background values, so they could not be analyzed the same way. Unfortunately, the concentrated assay was not able to be repeated multiple times, because it was carried out using the last of the remaining reagents from the HK activity assay kit.

The results of the HK activity assay of the concentrated lysate samples shows a higher HK activity in the wild-type sample than in the LPL-KD sample, which supports the initial HK activity assay results. Although this result was not what we were initially expecting, it does have several logical explanations. The first possible contributing factor

to this phenomenon lies with the concentrations of our two metabolic enzymes of interest,

HK II and PFK-1, in the two cell lines. As was previously stated, the LPL-KD cell line has significantly increased expression and translation of HK II, and a slight decrease in the expression of PFKM. The increased level of HK II in the cells means that more glucose can be utilized by the knock-down cells, but the decrease in PFK-1 suggests that there is further regulation preventing all of that glucose from being used in glycolysis.33

As a result, it is likely that the product of HK II, G6-P, would build up in the cells. Since this product is an inhibitor of the HK II enzyme, it is possible that this increased G6-P concentration has a stronger inhibitory effect on the HK II enzyme activity in the LPL-

KD cells than the LCACoAs have on the HK II enzyme activity in the WT cells.

Therefore, despite the fact that there may be more inhibition by fatty acid products in the

WT cells, it does not outweigh the inhibition caused by the buildup of G6-P in the LPL-

KD cells.

A second possible explanation for why the HK activity is higher in the WT cells comes from the conditions of cellular differentiation that were used to prepare our cells for lysis and analysis. The primary source of FFAs for the growing L6 cells is the fetal bovine serum used in the growth medium. The FBS most likely has VLDL particles (as reported on the FBS product datasheet), which are substrates for LPL action. The initial growth medium contains 10% FBS, and thus the growing and dividing cells have access to sufficient quantities of FFAs. However, the differentiation conditions utilize medium that has reduced FBS, with incubation in a 2% FBS medium for 2 days followed by incubation in a 0% FBS medium for 2 more days. It is possible that the 2 days of incubation in complete DMEM that contains 0% FBS gives the L6 cells sufficient time to

finish metabolism of the FFA derivatives that have been taken into the cells. If there is no more FFA product within the cells, then there would be no difference in inhibition by

FFAs in the WT cells compared to the LPL-KD cells, even though transcript and protein levels may remain altered several days after removal of FBS. It is note-worthy that under the same experimental conditions, LPL-KD cells demonstrated a greater glucose utilization (oxidation to CO2 and incorporation into glycogen).

Section 4 – Literature review of altered LPL expression and metabolism

Overexpression of LPL

The role of lipoprotein lipase in glucose utilization and insulin sensitivity has been a subject of research for many years. Kim et al. determined that mice with a 4-fold muscle specific upregulation of LPL (muscle-LPL mice) exhibited muscle triglyceride content that was three times higher than that of the control mice.34 The muscle-LPL mice also showed a 46% decrease in whole-body glucose uptake, as well as a 29% decrease in insulin-stimulated whole-body glycolysis. These findings provide support for the inverse correlation between LPL expression and cellular glucose utilization seen in our cellular experiments, and seem to have a likely biological cause. The muscle cells of the muscle-

LPL mice are higher in triglyceride and fatty acid content, so they are more readily able to obtain energy through the preferred pathway of fatty acid oxidation.4 For this reason, they would not require as much glycolysis, and thus would benefit less from insulin- stimulated glucose uptake.

The proposed mechanism of insulin resistance in the study by Kim et al. was a

63% decrease in insulin-stimulated activation of insulin receptor substrate-1 (IRS-1)-

associated phosphatidylinositol (PI) 3-kinase.34 IRS-1-associated PI 3-kinase activity has been correlated with insulin stimulation of glucose transport as well as glycogen synthase activity in skeletal muscle.35 This kinase is important in intracellular mediation of insulin signaling in skeletal muscle, so the decrease in muscle insulin response could be a result of the decreased insulin signaling in the muscle-LPL mice.34 However, the insulin resistance of the muscle-LPL mice was specific to the muscle tissue, as the insulin response in the liver remained unchanged. The possible regulation of IRS-1 and PI 3- kinase by the silencing of LPL in L6 skeletal muscle cells is being explored further by another student in our lab group.

LPL down-regulation in cardiac muscle

The impact of down-regulating LPL activity has also been studied in cardiac muscle. Cardiac muscle is similar to skeletal muscle because it utilizes fatty acid metabolism as its primary energy source, while still utilizing glucose and ketone bodies when needed.4 However, cardiac muscle does not undergo any anaerobic metabolism, so lactic acid fermentation never occurs. Augustus et al. investigated the metabolic effects of losing lipoprotein lipase-derived fatty acids on cardiac muscle through the study of heart-specific LPL knock-out mice (hLPL0).

LPL-derived FFAs are the primary fatty acid source for the heart, but cardiac muscle cells are also able to obtain fatty acids that are bound to albumin.36 Augustus et al. wanted to test the extent to which the two fatty acid sources are interchangeable. To do so, they studied the metabolic changes that occurred in the hLPL0 mice compared to wild-type mice. The absence of heart LPL led to an increase in circulating plasma triglycerides in the hLPL0 mice.37 Unsurprisingly, the mice which did not produce heart

LPL and had increased TG content showed a significant decrease in myocardial fatty acid oxidation. These hLPL0 mice also exhibited an increase in both myocardial glycolysis and glucose oxidation compared to their WT counterparts. Ultimately, it was determined that the fatty acids that can be obtained from serum albumin were not sufficient to replace those that come from cardiac muscle LPL activity. The hLPL0 hearts showed decreased ability to respond to aortic constriction and increased contractile dysfunction and cardiac fibrosis.37

The study of cardiac muscle in the absence of LPL further supports the data that has been observed by our lab. It showed that glucose oxidation in both LPL-KD cells and hLPL0 mice is up-regulated in order to make up for the decreased potential for energy production via fatty acid oxidation. Although Augustus et al. did not specifically look into the glycolytic gene expression, it is likely that they would have observed similar trends to those that we have seen in our skeletal muscle cells.

Chapter 5 – Conclusion

In summary, we were able to determine that the gene expression of PFKM was reduced in LPL-KD L6 cells, while the gene expression and protein concentration of HK

II was increased. The activity of hexokinase is higher in the wild-type cells, despite the fact that there is less enzyme produced. Our proposed hypothesis that explains this phenomenon is outlined in Figure 18.

Figure 18. Proposed explanation for the regulation of hexokinase II activity in LPL-KD cells. Our research further investigated the relationship between LPL expression, FFA availability, and glucose metabolism. Previous studies have shown that there is an inverse correlation between fatty acid metabolism and glucose metabolism, but the interplay between the two metabolic pathways and their associated enzymes are still not fully understood.38 This investigation into the impact of reducing LPL expression on the enzymes HK II and PFK-1 may help further the understanding of this complicated interaction.

We would continue this research by working on perfecting our analysis of the HK activity in the two cell types. Identifying the right amount of lysate to add to reduce baseline NADH production would greatly improve the ability of the assay to produce reliable results. Additionally, it would be beneficial to work out a way to treat the cells that will be used for the HK activity assays with chylomicrons, VLDLs, or FFAs in order to make sure that the inhibition by LPL products can be observed and compared between the different cell types.

We would also benefit from further research into the PFK-1 enzyme. It would be worthwhile to attempt the PFK-1 western blot with different antibodies, or different concentrations of antibodies, in order to get a good idea about the concentration of the enzyme within the LPL-KD and WT cells. Ideally, an activity assay that looks at PFK-1 activity would round out the research, and would help explain the HK activity data that was collected for this thesis.

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