THE EFFECT OF HIGH HYDROSTATIC PRESSURE ON STABILITY OF PYRUVATE

OXIDASE FROM AEROCCOCUS SPECIES

by

Luke Wallace

(Under the Direction of José Reyes I. De Corcuera)

ABSTRACT

Pyruvate oxidase (POX) catalyzes the oxidative decarboxylation of pyruvate to

acetylphosphate in the presence of oxygen, a divalent cation, and its cofactors thiamine

pyrophosphate and flavin adenine dinucleotide. High hydrostatic pressure (HHP) was applied at

selected temperatures to elucidate its effect on the thermostability of POX. High pressure

generally destabilizes POX. Rate of inactivation was slowed slightly between atmospheric

pressure and 50 MPa at 35 °C, where kinact decreased from 0.004 ± 0.003 min-1 to 0.001 ± 0.001

min-1. Activation volume halved to 29 ± 13 cm3/mol at 45 °C from 68 ± 43 cm/mol at 35 °C and

71 ± 12 cm/mol at 25 °C. Data from non-denaturing gel electrophoresis and ultraviolet

spectroscopy suggest that POX remains in a native conformation before dissociating and

precipitating with the application of HHP. Low density polyethylene reduced POX activity,

though the specific reason for this affect is unclear.

INDEX WORDS: Pyruvate oxidase; High hydrostatic pressure; Enzyme kinetics; Stabilization

THE EFFECT OF HIGH HYDROSTATIC PRESSURE ON STABILITY OF PYRUVATE

OXIDASE FROM AEROCCOCUS SPECIES

by

LUKE WALLACE

B.S.A., The University of Georgia, 2013

A Thesis Submitted to the Graduate Faculty of the University of Georgia in Partial Fulfillment of

the Requirements for the Degree

MASTER OF SCIENCE

ATHENS, GEORGIA

2017

© 2017

Luke Wallace

All Rights Reserved

THE EFFECT OF HIGH HYDROSTATIC PRESSURE ON STABILITY OF PYRUVATE

OXIDASE FROM AEROCCOCUS SPECIES

by

Luke Smith Wallace

Major Professor: José I. Reyes De Corcuera

Committee: Fanbin Kong

Derek Dee

Electronic Version Approved:

Suzanne Barbour

Dean of the Graduate School

The University of Georgia

August 2017

ACKNOWLEDGEMENTS

I would like to thank my major advisor, Dr. Reyes, for taking me on during a challenging

transition and for his constant support and enthusiasm throughout my project. I acknowledge and

appreciate the funding and financial assistance provided to me by Dr. Reyes, the department of

Food Science and Technology, and the USDA-NIFA-AFRI grant #2014-67021-21604, without

which my graduate education would not have been possible. To my advisory committee, Dr.

Kong and Dr. Dee, I thank you for your assistance and feedback in developing my project and

for your service in my committee. Lastly, I would especially like to thank my lab members, both

current and former, and my friends Dr. Garcia, Ali, Martina, Victoria, Daoyuan, Tristin, Brittnee,

Vivian, Hanna, Erica, Jess and many others, both for their help in my project, and for their

relentless support in helping me survive graduate school.

iv

v

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS.……………………………………………………...………………iv

LIST OF TABLES………………………………………………………………………………..vi

LIST OF FIGURES…………………………………………………………………………...…vii

CHAPTER

1. LITERATURE REVIEW……………………………………………………………..1

Introduction……………………………………………………………………………1

Characteristics of Pyruvate Oxidase…………………………………………………..2

Detection of Pyruvate………………………………………………………………....4

Applications for Pyruvate Oxidase in Biosensors…………………………………….6

High Hydrostatic Pressure Stabilization of Enzymes………………………………..13

Gap of Knowledge…………………………………………………………………...17

Hypothesis……………………………………………………………………………17

Objectives……………………………………………………………………………17

2. THE EFFECT OF HIGH HYDROSTATIC PRESSURE ON STABILITY OF

PYRUVATE OXIDASE FROM AEROCOCCUS SPECIES……………………….19

Introduction…………………………………………………………………………..19

Materials and Methods……………………………………………………………….20

Results and Discussion………………………………………………………………34

3. FINAL COMMENTS………………………………………………………………..49

REFERENCES…………………………………………………………………………………..51

APPENDIX……………………………………………………………………………………....57

vi

LIST OF TABLES

Page

Table 1.1. Comparison of biosensors based on pyruvate oxidase………………………….……10

Table 1.2. Pyruvate oxidase biosensor stabilization technique comparison……...………….…..12

Table 2.1. Reaction cocktail composition for pyruvate oxidase activity assay....……………….25

Table 2.2. Residual activity for pyruvate oxidase treated using original pouch design at 100

MPa at 5 °C relative to both 0 s treatment time and fresh working enzyme solution

(n=3)…………………………………………………………………………………...…30

Table 2.3. Residual pyruvate oxidase activity for samples sealed in new pouches relative to

fresh enzyme activity over time………………………………………………..………...31

Table 2.4. First order rate of pyruvate oxidase inactivation ± standard error determined by the

linear regression of residual activity versus treatment time….…………………………..39

Table 2.5. Residual activity data for confirmation experiments using new pyruvate oxidase

batch compared to residual activity for the old batch calculated based on kinact.………...42

Table 2.6 Apparent activation energy of inactivation ± standard error determined by the linear

regression of ln(k) versus 1/T……………...…………...……………………………….43

Table 2.7 . Activation volume ± standard error determined by the linear regression for each

temperature………………………………………...…………………………………….45

vii

LIST OF FIGURES

Page

Figure 1.1. Typical elliptical pressure-temperature diagram with active and inactive enzyme

regions……………………………………………………………………………….…..15

Figure 1.2. Monomeric subunit of pyruvate oxidase (PDB ID: 1VF5) with its cofactors FAD

(1) and TPP (2), three largest cavities (3-5), and active site, GLU54 (6), labeled.….…..16

Figure 2.1. High hydrostatic pressure system...……………………………………………….....22

Figure 2.2. Pressure (○) and temperature (□) profiles for a sample treated at 35 °C at 100 MPa

for 300 s and pressure (●) and temperature (■) profiles for a sample treated at 35 °C

at 100 MPa for 0 s ……………………………………………………….………………24

Figure 2.3. Comparison of original (left) and newer (right) pouch design to scale………...……29

Figure 2.4. Residual activity relative to fresh enzyme over time for pyruvate oxidase in low-

density polyethylene pouch ( ), high density polypropylene tube ( ), and amber glass

vial ( )..……….…….…………………………………………………………………....32

Figure 2.5. Corrected absorbance measurements for individual replicates of fresh pyruvate

oxidase (black) compared with samples treated at 0.1 MPa at 35 °C for 0 (dark grey)

or 600 s (light grey)…………………………………………………………………........34

Figure 2.6. Residual activity for t0 treatments relative to fresh enzyme at 25 ( ), 35 ( ), or

45 °C ( )…………………………….……………………………………………………35

Figure 2.7. Reactor (open symbol) versus syringe (closed symbol) temperature at 25 (light grey),

35 (dark grey), or 45 °C (black)……………………………………………………….....36

viii

Figure 2.8. Residual activity for POX at 25 (A), 35 (B), and 45 °C (C) at 0.1 ( ), 50 ( ),

100 ( ) or 150 ( ) MPa…………………………………………………………………..37

Figure 2.9. Logarithm of residual activity at 25 (A), 35 (B), and 45 °C (C) at 0.1 ( ), 50 ( ),

100 ( ) and 150 ( ) MPa………………………...……………………………………….40

Figure 2.10. Rate constant of inactivation versus pressure at 25 ( ), 35 ( ), and 45 °C ( )….….45

Figure 2.11. Relative distance traveled through 4-20 % polyacrylamide gel by native and

inactivated pyruvate oxidase samples ………………………………………………..….47

Figure 2.12. Ultraviolet spectra for Native (black), Inactivated (grey), and blank (dotted)

samples …………………………………………………………………………………..47

Figure A.1. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 0.1 MPa at 25 °C

for 3000 s………………………………………………………………………………...57

Figure A.2. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 50 MPa at 25 °C

for 0 s ……………………………………………………………………………….…...58

Figure A.3. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 50 MPa at 25 °C

for 3000 s……………………………………………………………...…………………58

Figure A.4. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 0.1 MPa at 35 °C

for 0 s ………………………………………………………………………………...….58

ix

Figure A.5. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 0.1 MPa at 35 °C

for 3000 s …………………………………………………………………………...…...59

Figure A.6. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 50 MPa at 35 °C

for 0 s …………………………………………………………………………………....59

Figure A.7. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 50 MPa at 35 °C

for 3000 s …………………………………………………………………………...…...59

Figure A.8. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 0.1 MPa at 45 °C

for 0 s ………………………………………………………………………………...….60

Figure A.9. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 0.1 MPa at 45 °C

for 1000 s ……………………………………………………………………….…...…..60

Figure A.10. Pressure (closed symbol) and temperature (open symbol) for samples one

(dark grey) and two (light gray) for supplemental sample treated at 50 MPa at 45 °C

for 0 s ……………………………………………………………………...………….…60

1

CHAPTER 1

LITERATURE REVIEW

Introduction

Pyruvate oxidase (POX) catalyzes the oxidative decarboxylation of pyruvate into

acetylphosphate and hydrogen peroxide in the presence of oxygen and phosphate. Pyruvate

oxidase is a homotetrameric flavoprotein that is integral to the process of microbial aerobic

respiration. The high substrate specificity of POX enables it to be utilized in electrochemical

biosensors used in the quantitative analysis of both pyruvate and phosphate.

Biosensors are useful tools for detecting or quantifying the concentration of specific

chemicals or biological materials. Because they offer high sensitivity and specificity while

maintaining short response times, simple analysis methods, and relatively low costs, biosensors

are often preferable to other analytical methods. Biosensors using POX have been constructed

both for the detection of pyruvate and phosphate. In the case of phosphate detection, biosensors

are preferable to conventional spectrophotometric methods due to faster measurement speed,

reusability, higher specificity, and the potential for miniaturization and transportability [1, 2]. In

the case of pyruvate, biosensors can achieve a lower limit of detection, potentially at a lower cost

than many conventional detection methods [3, 4].

Pyruvate oxidase biosensors have potentially important roles in the food industry, the

medical field, and in environmental science. However, due to the poor stability of POX, current

2

biosensors lack the longevity to make them practical tools for most applications. Table 1.1

compares existing POX biosensors with respect to their operational and storage stability.

High hydrostatic pressure (HHP) has a stabilizing effect on many enzymes, and in certain

cases has been shown to enhance enzymatic activity [5]. It is not yet clear if HHP alone can be

used to enhance the practicality of enzymatic biosensors, but it could play a synergistic role

when paired with other stabilization techniques such as chemical, physical, and/or genetic

modification towards creating a more stable biosensor.

This chapter will review the current knowledge of POX activity, stability, and stabilization

within the context of biosensor fabrication.

Characteristics of Pyruvate Oxidase

Pyruvate oxidase is a homotetrameric flavoprotein in which each unit (65.5 kDa) binds

one FAD and one thiamine pyrophosphate (TPP) in the presence of a divalent cation, most

commonly Mn2+ or Mg2+. Pyruvate oxidase catalyzes the oxidative decarboxylation of pyruvate

according to the equation:

𝐶𝐻3𝐶𝑂𝐶𝑂𝑂− + 𝑂2 + 𝐻𝑃𝑂4

−2 + 2𝐻+ → 𝐶𝑂2 + 𝐻2𝑂2 + 𝐶𝐻3𝐶𝑂𝑂𝑃𝑂3𝐻−

The catalytic activity of POX can be broken down into five steps. The first is the deprotonation

of the C2-H of TPP, as with all TPP-dependent enzymes. Secondly, pyruvate is bound to the C2

atom of enzyme-bound TPP. Next is the decarboxylation of pyruvate to form hydroxyethyl-TPP,

followed by the oxidation of hydroxyethyl-TPP by FAD, and finally the reoxidation of FAD by

oxygen [6].

The optimum temperature for POX from Aerococcus sp. is 25 °C [7], 38 °C from

Lactobacillus plantarum [8], and 42 °C from Escherichia coli [9]. The optimum pH for POX

from Aerococcus is 7 [1] and 5.6 from Lactobacillus plantarum [8].

3

Size-exclusion HPLC and analytical ultracentrifugation show that the holoenzyme retains

its tetrameric state down to 20 µg/mL. The apoenzyme, however, shows a stepwise tetramer-

dimer-monomer dissociation at a protein concentration of 20 µg/mL. The quaternary structure is

stabilized in the presence of the cofactors, TPP, FAD, and a divalent cation. In the presence of

divalent cations, FAD, and TPP bind to the apoenzyme, forming the binary complex. Both TPP

and FAD affect the association of the quaternary structure by shifting the equilibrium towards a

dimer or tetramer structure. High FAD concentrations elicit a strong stabilizing effect against

urea and heat denaturation, while excess TPP has no effect [10].

POX activity requires the presence of a divalent metal cation. Blake, et al. [11] compared

the activity of pyruvate oxidase in the presence of Mg2+, Ca2+, Zn2+, Mn2+, Ba2+, Ni2+, Co2+,

Cu2+, and Cr3+. The results showed that while the most consistent steady-state kinetics were

observed with Mg2+ and Mn2+, the enzyme lacked specificity for the metal ion required for

catalysis. These findings suggest that the divalent metal cation is not a true cofactor of POX, but

that the cation instead binds to TPP, forming a metal-TPP complex that is the true enzymatic

cofactor.

To elucidate the conditions for thermal inactivation of the enzyme, POX from Bacillus

stearothermophilus was isolated along with other enzymes such as cytochrome oxidase, malic

dehydrogenase, and adolase from the cell granule using lysozyme. Unlike all other enzymes

isolated from the thermophile in this study, POX was quickly inactivated at the optimal growing

temperature range for the bacteria (60-65 °C). After one hour at 60 °C, the activity of POX

decreased to only 12%. The presence of pyruvate, Mg2+, and oxygen all increased thermal

stability. Pyruvate oxidase left within unlysed cells showed stability at 60 °C up to an hour, at

which point the system began to run out of oxygen and substrate [12].

4

Chang and Ruan [13] explored the pressure-induced dissociation of pyruvate oxidase

from E. coli using a fluorescence spectrophotometer attached to a pressurized chamber to

monitor the enzyme’s dissociation from 0.1 to 300 MPa. Results indicated that the native enzyme

dissociated completely at 220 MPa, but that the dissociation was likely reversible. FAD had a

stabilizing effect against pressure. When FAD was removed from the native enzyme, the

resulting apoenzyme had a half-dissociation pressure of only 98 MPa as opposed to 130 MPa for

the native enzyme. Based on the fluorometric data, the dissociation of the apo-pyruvate oxidase

occurred in a single step, the dissociation of the four subunits. In contrast, the native enzyme

showed an extra step wherein FAD was removed before the full dissociation of the subunits

occurred. This, along with calculated changes in molar volume, led to the author’s theory that

FAD induces a conformation change resulting in better matching of the subunit interface within

POX. Lastly, as enzyme dissociation was compared at pH values of 7.5, 8.5, or 9.5, results

showed that stronger alkalinity promoted POX dissociation. This may be due to a decrease in

subunit affinity caused by a change in ionization of amino acid sidechains. In the case of acidic

pH values, measurements were impossible due to enzyme precipitation under high pressure.

Detection of Pyruvate

Pyruvate is an important part of many metabolic pathways, perhaps most notably as a

product of glycolysis and as a starting point for the Krebs cycle, oxidative phosphorylation, and

amino acid synthesis. The detection of pyruvate is important to the food industry, the medical

field, and in environmental science.

The detection of pyruvate is of importance to the process of food fermentation [14].

During the production of wine and beer, the presence of pyruvic acid and other keto acids in the

beer wort or grape must is associated with important flavor compound production by yeast in the

5

finished product. During fermentation, specific amino acids (valine, leucine, isoleucine,

methionine, and phenylalinine) are metabolized by Saccharomyces cerevisiae into keto acids that

cannot be used in primary carbon metabolism. These acids are redirected into the Ehrlich

pathway where they are converted into fusel alcohols, which unlike ethanol contain more than

two carbons. In high concentrations, fusel alcohols contribute undesirable off-flavor. However,

in lower, controlled concentrations, these products help to contribute to the distinct flavor

profiles of many products [15, 16].

In onions, pyruvate serves as an important quality indicator of pungency [17].

Schwimmer and Weston [18] suggested that pungency arises from an interaction of L-cysteine

derivatives and allinase enzymes when the onion tissue is damaged, resulting in the production

of pyruvic acid. Biosensors made from immobilized pyruvate oxidase have differentiated

between low and high-pungency onion by measuring pyruvate concentrations [4]. Conventional

pyruvate measurements for alliums can be time consuming and expensive. With low-pungency

onions growing in popularity, means of accurately measuring pyruvate are becoming more

important [19].

Pyruvate oxidase biosensors have potential applications in the medical field. The

monitoring of L-lactate dehydrogenase (LDH) in human blood serum is important for medical

analysis. It catalyzes the reaction:

𝐿 − 𝑙𝑎𝑐𝑡𝑎𝑡𝑒 + 𝑁𝐴𝐷+𝐿𝐷𝐻→ 𝑝𝑦𝑟𝑢𝑣𝑎𝑡𝑒 + 𝑁𝐴𝐷𝐻 + 𝐻+

The conversion of NAD+ to NADH can be measured spectrophotometrically, but this is

difficult to do in colored samples, such as blood serum. Pyruvate oxidase presents a potential

alternative to the rapid detection of LDH in serum [20]. Additionally, the monitoring of

pyruvate in blood serum as fluctuations can be indicative of diabetic acidosis or vitamin B1

6

deficiency [14]. Similarly to LDH, pyruvate in serum can be monitored spectrophotometrically,

but problems arise due to the turbidity of blood samples.

Perhaps the main application for pyruvate oxidase-based biosensors is the monitoring of

phosphorous in aquatic environments. Phosphorous is an essential component to metabolism in

all known life forms, serving as an essential building block for nucleic acids and various

intermediate metabolites [21]. Monitoring phosphorous levels in lakes and rivers is a major

environmental concern due to eutrophication. Eutrophication occurs when waters become over-

enriched with nutrients, with phosphorous being the most common cause of eutrophication in

freshwater. During eutrophication, the over-enrichment of nutrients in water increases the

production of autotrophs such as algae and cyanobacteria. The accompanying increase in cellular

respiration causes hypoxia or anoxia, resulting in the death of aquatic animals and a detriment to

biodiversity. The detection of phosphorous is particularly important, because unlike nitrogen and

carbon which can be acquired in the atmosphere, phosphorous is usually found in more limiting

amounts, most commonly from surface water runoff. Human activities, mainly agriculture, are

the main contributors to phosphorous in surface runoff [22].

Applications for Pyruvate Oxidase for Biosensors

Existing alternatives for Pyruvate and Phosphate Detection

A biosensor is an integrated receptor-transducer device that converts a biologically-

induced recognition event, such as enzyme activity, into a detectable signal via a transducer and

processor. The signal can be used to depict the presence and/or concentration of the analyte of

interest using a display. Electrochemical biosensors are based on the monitoring of either the

production or consumption of electroactive species. Transduction occurs under a variety of

methods that fall under either potentiometry or amperometry. Potentiometric biosensors monitor

7

the electric potential of an electrochemical cell in a working electrode relative to a reference

electrode with zero current flow. Measuring the concentration of the target analyte can be

achieved by measuring ions or gases that are either generated or consumed during enzymatic

activity. Amperometric biosensors tend to be more common, both for single-use and multi-

measurement devices. Unlike potentiometric devices, amperometric biosensors utilize a constant

potential applied between a working and a reference electrode. The resulting net current flow

from the imposed potential is proportional to the concentration of the electroactive species in

solution [23].

Akyilmaz and Yorganci [24], developed a biosensor to investigate the effect of thiamine

on pyruvate oxidase. The biosensor was prepared by covalently immobilizing pyruvate oxidase

onto a dissolved oxygen probe using gelatin and glutaraldehyde as a cross-linking agent. During

the enzymatic reaction, the dissolved oxygen layer in the interval space decreased in relation to

concentration of added substrate. The difference in dissolved oxygen (ΔDO) was measured

between before and after the addition of substrate to the reaction medium. The principle of the

biosensor relied on detecting this ΔDO in relation to thiamine concentration to construct a

standard curve for the determination of thiamine. All measurements were done at 35°C in a

thermostatic reaction cell filled with oxygen-saturated phosphate buffer (50 mM, pH 7.0). The

biosensor could detect a linear range of thiamine over a constant concentration of pyruvate.

As previously mentioned, while phosphorus is an essential element to all life, elevated

levels caused by surface runoff from agricultural or other human activities can cause an

excessive growth of algae or cyanobacteria, leading to eutrophication [25]. Pyruvate oxidase is

the most commonly-used enzyme for multi-enzymatic phosphate biosensors. These are generally

considered to be simpler to construct and operate than other multi-enzyme phosphate biosensors

8

[26]. Gavalas and Chaniotakis [27] created a phosphate biosensor using recombinant pyruvate

oxidase from Lactobacillus plantarum stabilized by a polyelectrolyte on a porous carbon rod.

Two polyelectrolytes were tested, diethylamonoethyl-dextran (DEAE-Dextran) at concentrations

0.10, 0.25, and 0.50 (%w/v) and DNA at 0.10 and 0.25 (%w/v) the 0.10 [DEAE-Dextran} and

0.25 [DNA] were chosen for further analysis, as they had the greatest sensitivity and stability of

those polyelectrolyte concentrations tested. The DEAE-Dextran showed the greatest stability,

with 67% remaining activity after 220 hours, compared with 56% for DNA, and 50% for the

control with no polyelectrolyte. Ogabiela, et al. [28] developed a highly-sensitive (limit of

detection o.1 µM) phosphate biosensor for use in freshwater sources. Using a highly-ordered

gold nanowire array (AuNWA) that integrated the enzyme’s cofactors, the concentration of POX

required could be reduced around 100-fold when compared to similar biosensors. The AuNWA

biosensor was resistant to interference from common freshwater interferants such as chloride,

sulfate, fluoride, nitrite, and nitrate ions. The device showed no loss in activity after two weeks

of daily measurements, and with a recovery of 96.6 ± 4.9%, making it one of the most stable and

most sensitive POX-based biosensors designed for detecting phosphate in water sources.

Pyruvate oxidase biosensors have low operational and storage stability compared to other

enzymatic biosensors. This is due in most part to the poor stability of POX [29]. Currently, many

techniques have been explored aiming to stabilize POX for use in biosensors. Table 1.2

compares different POX stabilization techniques in biosensors. The stability of POX biosensors

has seen only moderate improvement since one of the earliest designs by Zapatabacri and

Burstein [14] in 1987. However, the sensitivity of biosensors based on POX has improved over

the past few decades. One of the most recent devices, the previously-mentioned design by

Akyilmaz and Yorganci [30] in 2007 using POX from Aerococcus sp., has produced the most

9

sensitive pyruvate biosensor to date, capable of detecting pyruvate linearly between 0.0025 –

0.05 µM, while still contending with other designs in terms of stability.

10

Table 1.1. Comparison of biosensors based on pyruvate oxidase

Substrate

Detected

POX

Source

Immobilization

Technique

Working

Electrode

Detection

Limit

Linear

Range

Operatio

nal

Stability

Storage

Stability

Ref Notes

Pyruvate Pediococ

cus spp.

Physical adsorption

with medolas blue

dye

Screen-

printed,

medolas

blue

mediated

electrode

1-2

µmol/g

fresh

weight

1-4

µmol/g

fresh

weight

[4] Disposable

biosensor to

detect pyruvate

in onions

Pyruvate Aerococc

us sp.

Cross-linked with

gelatin using

gluteraldehyde

YSI type

DO probe

covered

with

Teflon

membrane

0.0025

-

0.05µM

86% after

9 hours

[30]

Pyruvate Aerococc

us sp.

Cross-linked with

gelatin using

gluteraldehyde

YSI type

DO probe

covered

with

Teflon

membrane

0.0025

-0.05

µM

90% after

20 days

[24] Investigating the

effect of

thiamine on

POX

Pyruvate Electropolymerizati

on, conductive

redox polymer

Glassy

carbon

electrode

1 µM –

1.8 mM

[31]

Pyruvate Recombi

nant POX

from

Lactobac

illus

plantaru

m

Co-immobilized

with horseradish

peroxidase in a

carbon paste using

methylene green

Modified

carbon

paste

electrodes

0.1-3

mM

[32]

11

Pyruvate Microorg

anisms

Modified pyrrole

monomer with

thiophene

Platinized

glassy-

carbon

electrode

0.0 –

1.0 mM

[29] Attempting to

develop

biosensor

capable of

detecting in O2-

free samples

Phosphate Recombi

nant POX

from

Lactobac

illus

plantaru

m

Genetic

modification and

physical adsorption

onto the carbon

tube

Porous

carbon

soaked in

polyelectr

olyte

solution

<0.3 mM 0.05 –

1.0 mM

67% after

220 hours

49% after

24 weeks

[27] Optimized to

measure

phosphate ion

activity in serum

Pyruvate 34 µM 90 –

600 µM

[33] Measured

pyruvate in

onions and garlic

Pyruvate Chemically bound

using methylene

green

Modified

carbon

electrode

0.38 –

1.03

mM

[34]

Pyruvate Chemically bound

to olyazetidine

prepolymer, nylon

membrane,

Modified

carbon

electrode

1-10

mM

[35]

Phosphate Aerococc

us sp.

Immobilized on

polypyrrole

Polished

platinum

3 µM 15-400

µM

[2]

Pyruvate Covalent

attachment to

polytyramine

Glassy

carbon

electrode

0.05 µM 0.1 –

3.0 mM

74% after

50 days

[3]

Pyruvate Cross-linked with

gelatin using

gluteraldehyde

pO2 meter Up to 2

mM

“several

months”,

~250

assays

[14]

12

Table 1.2. Pyruvate oxidase biosensor stabilization technique comparison

Method Most Stable Version Measure of Stability Ref.

POX immobilized with gelatin and

insoluble film using glutaraldehyde

(GA) crosslinked with glucose

35 °C, pH 7.0, 50

mM phosphate buffer

>85% activity

remaining after 9

hours at 35 °C

[30]

Electropolymerization of

mercaptohydroquinone in the presence

of POX

0.2 mM TPP, 10 µM

MgSO4 at 20 °C

5% loss after 7 days

stored at 5 °C

[31]

POX immobilized using GA on a self-

assembled monolayer made of 3-

mercaptopropionic acid and 6-

aminocaproic acid

4-hour assembly of

self-assembled

monolayer

90.5% activity after

15 assays, 26% loss

after 10 days of

refrigerated storage

[36]

Recombinant POX from Lactobacillus

plantarum co-immobilized with

horseradish peroxidase in organic carbon

paste

Addition of raffinose

and protective

dialysis membrane

89% after 3 months,

66% after 6 months

with storage in

desiccator at 4 °C

[32]

POX from Aerococcus sp. crosslinked

on Pt/Au alloy nanowires along with

cofactors

40%Pt + 60% Au,

150 mM TPP, 5 µM

FAD, 10 U/L POX

48% loss after 2

weeks, stored in 0.1

M citrate buffer, pH

7.0 at 4 °C

[37]

POX from L. plantarum covalently

immobilized on copolymer poly (5-

hydroxy-1,4-napthoquinone-1,4-

nathoquinone acid)

pH 7.5 40% loss after 3 uses,

50% loss after 7 days

in storage at 4 °C

[38]

Potentiostatic redox film with

polypyrrole film to prevent POX

leaching

5mM, oxygen-

saturated phosphate

buffer on pure

polythiophene film

14% remaining after

10 days stored in pH

6.5 phosphate buffer

along with cofactors

at 4 °C

[29]

Recombinant POX from L. plantarum

stabilized with polyelectrolyte DEAE-

dextran or DNA

0.10% w/v DEAE-

dextran with 20-hour

polarization

67% after 220h, 49%

after 24 weeks under

dry storage at -20 °C

[27]

POX from Aerococcus viridans

immobilized by nafion matrix covered in

poly (caromoyl) sulfunate hydrogel

Use of both nafion

and PCS hydrogel

>85% after 12 hours

of continuous

operation

[1]

POX from Aerococus sp. crosslinked

with bovine serum albumin and

glutaraldehyde along with cofactors on

gold nanowire array

Crosslinkage with

both BSA and GA

100% activity after 2

weeks of repeated

use, stored in 50Mm

citrate buffer in

refrigerator

[28]

13

High Hydrostatic Pressure Stabilization of Enzymes

The application of high hydrostatic pressure (HPP) is effective in inactivating certain

deteriorative enzymes in foods [39, 40]. Pressure has often been viewed as a protein denaturant,

as it disrupts the multimeric structure of many proteins. There exists, however, examples of

pressure-enhanced stability in proteins, due mainly to pressure’s effect on hydrophobic, van der

Waals, and electrostatic interactions [41]. Generally, the effect of pressure on physiochemical

processes at equilibrium are governed by Eyring’s Equation [42]:

(𝛿 ln 𝑘

𝛿𝑝)𝑇 = −(

Δ𝑉≠

𝑅𝑇)

where k is the rate constant, p is pressure, T is absolute temperature, R is the ideal gas constant,

and ΔV≠ is the activation volume that represents the influence of pressure on the reaction rate.

This equation can be integrated and rearranged to produce the equation:

ln 𝑘 = −∆𝑉≠

𝑅𝑇×𝑃 + ln 𝑘𝑝0

where k is the rate constant at a reference pressure, p0. This means that a reaction with a

negative volume change will be shifted towards the more compact state under pressure, while

one with a positive volume change will be slowed.

For changes in protein structure, the precise magnitude and sign of ΔV≠ is dependent

upon the specific molecular interactions [43].. Furthermore, unlike chemical catalysis, calculating

ΔV≠ can be difficult to calculate for enzymes under pressure, as it is dependent on changes in

enzyme conformation, enzyme solvation (including interactions with media and other proteins),

specific chemical equilibrium, and interactions with other proteins. Values for ΔV≠ typically

have a magnitude of less than 30 cm-3mol-3, but values can range from -70 to +60 cm-3mol-3 [44].

14

Noncovalent interactions are the primary molecular force altered through pressure. The

solvation of charged groups is associated with a reduction in volume, whereas the formation of

coulombic interactions accompanying the dehydration of charged atoms results in an increase in

volume. The formation of hydrophobic interactions between aliphatic groups (associated with a

positive ΔV≠) is destabilized under pressure, whereas the stacking of aromatic rings is favored by

increased pressure, as it is associated with a small decrease in volume. Hydrogen bonding is

associated with virtually no change in volume, and is therefore largely independent of pressure

[45, 46]. Under a high-pressure simulation of bovine pancreatic trypsin inhibitor (BPTI),

Kitchen, et al. [47] found that the hydration shell around the molecule was more ordered under

high pressure as compared to the low pressure simulation. This effect was especially pronounced

around more nonpolar surface groups.

In studying the stabilizing effect of pressure on α-chymotrypsin against thermal

inactivation, Mozhaev, et al. [48] suggested that the stabilizing effect of pressure may be due to

the opposing effects of temperature and pressure on the ability of protein functional groups to

interact with water. This was supported by Taniguchi and Suzuki [49], who stated that while

increased heat is accompanied by protein unfolding, the resulting unfolding exposes more

hydrophobic residues to the surrounding water. Under high-pressure conditions, the hydration of

these newly-exposed functional groups is associated with a decrease in molar volume of water

with the increase in ordering of the hydration shell [50]. It is likely that the promotion of

nonpolar functional groups interacting with water under high pressure is key to the stabilizing

effect of pressure against thermal inactivation. Hawley [51] devised an elliptical model (Figure

1.1) to describe the effects of pressure and temperature on enzyme conformation. The active

conformation is represented by the darker, inner region, surrounded by the lighter region

15

representing the point of reversible denaturation, with all areas outside the ellipse representing

complete denaturation.

Figure 1.1. Typical elliptical pressure-temperature diagram with active and inactive enzyme

regions

More recent studies have examined the role that hydrophobic cavities play in the

behavior of proteins under HHP. The decrease in volume associated with protein unfolding under

pressure has been attributed to the difference between the density of bulk water and water

associated with the protein, the loss of internal cavities, or some combination of the two. Roche,

et al. [52] studied the structural and energetic details of unfolding in several staphylococcal

nuclease mutants with varying cavity sizes using simulations based on high-pressure NMR

spectroscopy. They determined that greater internal void volume increased the magnitude of the

volume change associated with unfolding, and that internal cavities were the primary

determinants of pressure unfolding in proteins. While the hydrophobic effect would predict that

the transfer of nonpolar hydrocarbons from the interior of the protein into water would be

expected to be unfavorable [53], Collins, et al. [54] found that, at high pressures, the filling of

these hydrophobic cavities became so favorable that new cavities would grow to accommodate

water molecules. In another homotetrameric oxidoreductase, urate oxidase, the high-pressure

Pressure

Active/Native

Inactive/Denatured

Reversibly Denatured

Tem

pera

ture

16

perturbation of a large hydrophobic cavity near the active site led to the unfolding of that enzyme

until quaternary structure damage caused irreversible aggregation [55]. In POX, there are three

cavities large enough to accommodate at least one water molecule (≥ 30 Å3) located directly

beside the active site, around GLU54 in each of its four subunits. Figure 1.2 shows a rendering of

one subunit of POX from Aerococcus viridans generated from crystallographic data from Juan,

et al. [56], with its three largest cavities and active site labeled.

Figure 1.2. Monomeric subunit of pyruvate oxidase (PDB ID: 1VF5) with its cofactors FAD (1)

and TPP (2), three largest cavities (3-5), and active site, GLU54 (6), labeled.

1

2

3

4 5

6

17

Gap of Knowledge

Many studies have utilized HHP to stabilize enzymes. While the exact mechanisms by

which HHP increases thermal stability in enzymes are unknown, it is thought that the effects of

HHP and temperature counteract each other in certain cases. Specifically, greater pressure

increases order, thereby decreasing entropy, while increased temperature decreases order, thus

increasing entropy [57]. Newer studies have suggested that the size and location of internal

cavities in the enzyme play an important role in how that protein behaves under HHP [52, 54,

55]. There exists no current mathematical model for predicting the effect of HHP on a given

enzyme based on molecular structure. Empirical models must be constructed for each enzyme to

observe the effects of HHP. At this point, no such empirical model exists for POX.

In our laboratory, we recently studied the effect of HHP on glucose oxidase[58], xanthine

oxidase [59], and alcohol oxidase [60]. These enzymes were stabilized under HHP, but the effect

of HHP on POX from Aeroccus sp. is unknown. Though HHP has been used to stabilize

enzymes, stability does not persist after depressurization. If there is a rapid loss in stability for

the presumptive HHP-stabilized POX, techniques would need to be developed to capture the

stabilized form onto the working electrode. This could potentially mean constructing the POX-

based biosensor under high pressure conditions. Further experiments would be required to

determine if the immobilized HHP-stabilized POX biosensor offered improvements in stability,

sensitivity, and reproducibility over currently POX biosensor models.

18

To characterize the effect of HHP on POX and determine if there is an optimal pressure-

temperature combination for maximal stability.

Hypothesis:

High hydrostatic pressure will stabilize pyruvate oxidase

Overall Objective:

19

CHAPTER 2

THE INFLUENCE OF HIGH HYDROSTATIC PRESSURE ON STABILITY OF PYRUVATE

OXIDASE

Introduction

The rapid and accurate detection and quantification of pyruvate and phosphorous is

important to the food industry, and for the fields of medical and environmental science. For

flavor and quality purposes, it is important to monitor the starting pyruvate levels in fermented

products such as beer wort or wine must [15]. Pyruvate levels in onions are a predictor of

pungency, which is an important indicator of quality for that crop [4, 19]. Detection of pyruvate

in blood serum can assist in clinical diagnoses, as fluctuations in pyruvate levels in blood serum

can be indicative of a variety of medical issues [14]. Accurate on-site monitoring of phosphorous

levels in freshwater is vital for predicting and controlling the environmentally-destructive

process of eutrophication [22]. Current analytical methods mainly involve spectroscopy, which

currently lack the transportability and specificity needed for on-site environmental measurements

of phosphorous, the specificity and ease of use for pyruvate in food samples, and may not be able

to measure turbid blood samples [1, 4, 20].

Pyruvate oxidase (POX) is a homotetrameric flavoprotein with a molecular weight of 262

kDa. It catalyzes the oxidative decarboxylation of pyruvate to acetylphosphate (Equation 2.1) in

the presence of oxygen. Current designs for POX-based biosensors offer higher specificity and

20

sensitivity than other analytical methods for pyruvate and phosphorous, but are limited due to the

poor thermal stability of the enzyme.

Originally, high hydrostatic pressure (HHP) was used as a nonthermal means of

inactivating deleterious enzymes in food, but more recently the utility of HHP has expanded.

Under certain combinations of pressure and temperature, many enzymes have shown enhanced

stability [5]. Presently, there exist no mathematical models for predicting the effect of pressure

on enzymes based on molecular structure, so we currently must rely on empirical data built on a

case-by-case basis for each enzyme.

Before POX biosensors can be effectively utilized, better enzyme stabilization techniques

need to be developed. Currently, POX stabilization has largely only been studied in the context

of constructing biosensors. The most common method of stabilization is by crosslinking the

enzyme, usually using glutaraldehyde [14, 24, 30]. To the best of our knowledge, no previous

study has examined the effect of HHP on the stability of POX. The objective of this research was

to examine the effect of HHP on POX at selected temperatures, and to determine if there is an

optimal pressure-temperature combination for promoting stability.

Materials and Methods

Materials and Equipment

Pyruvate oxidase from Aerococcus sp. (product number P4105-100UN), peroxidase from

horseradish (POD, product number P8250-25KU), flavin adenine dinucleotide (FAD), sodium

(2.1)

21

pyruvate, and thiamine pyrophosphate (TPP), N-Ethyl-N-(2-hydroxy-3-sulfopropyl)-m-toluidine

(EHSPT), ethylenediaminetetraacetic acid (EDTA), magnesium sulfate, and 4-aminoantipyrine

were all purchased from Sigma-Aldrich (St. Louis, MO, USA) for use in the enzymatic activity

assay for POX. Enzyme buffers were made from potassium phosphate disodium salt from Fisher

Scientific (Pittsburg, PA, USA). Activity was assayed spectrophotometrically using a BioTek

Synergy™ HTX Multi-Mode Microplate Reader (VT, USA) using a xenon flash lamp and

monochromator at 550 nm. Data was collected using BioTek’s Gen5 Data Analysis Software.

For treatment of POX samples, the HHP system (Figure 2.1), was composed of a

micropump (model MP5), an 8.5-mL high pressure reactor (model U111), and a pump controller

(MP5 micropump control unit), from Unipress Equipment (Warsaw, Poland). Reactor

temperature was controlled by water baths, Isotemp 3016D (5 °C) and Isotemp 6200 R28 (25 -

45 °C) from Fischer Scientific (Pittsburg, PA, USA), which fed water through the jacket of the

high-pressure reactor. Selection of flow from each water bath was controlled by a pair of Sirai

Z110A solenoid pinch valves (Busseri, Italy). Temperature was monitored by a type K

thermocouple inserted through the bottom of the reactor with the tip flush to the bottom of the

vessel. A program written in LabVIEW controlled process time, pressure, and the activation of

solenoid valves with a data acquisition board (NI cDAQ 9174) from National Instruments

(Austin Texas). Pressure, time, and temperature were recorded by the LabVIEW program.

Enzyme samples were treated in modified 0.5 mL high-density polypropylene syringes (Medi-

Dose, Ivyland, PA, USA) added to the reaction vessel with Sil 180 oil bath liquid (Thermo

Scientific, Rockford, IL, USA). This was the same system used by Eisenmenger and Reyes-De-

Corcuera [57], apart from an additional BK Precision 1666 DC regulated power supply (B&K

Precision, Yorba Linda, CA, USA) used to isolate on of the DAQ modules to rectify a ground

22

loop issue that was discovered to cause inaccurate temperature readings during preliminary

measurements.

Figure 2.1. High hydrostatic pressure system.

The native polyacrylamide gel electrophoresis (PAGE) system consisted of 4-20%

polyacrylamide pre-cast gels (product number 456-1094) placed inside a Mini-PROTEAN®

Tetra cell, powered by a Power Pac 3000 power supply, all purchased form Bio-Rad Industries

(Hercules, CA, USA). Tris/glycine running buffer, native running buffer (product number 161-

0738), and Coomassie G-250 stain were also purchased from Bio-Rad. Staining occurred on a

Red-Rotor PR70-115V shaker platform (Hoefer Scientific, Holliston, MA, USA).

Sample spectra for native and inactivate POX samples were recorded using a

NanoDrop™ One Microvolume UV-Vis spectrophotometer (Thermo Fisher Scientific, Waltham,

MA, USA).

23

High Hydrostatic Pressure Processing

Syringes were modified by removing the plunger and truncating the body of the syringe

at the 0.15 mL mark before replacing and trimming the plunger. A 100 µL sample of POX was

slowly pipetted into a modified syringe while simultaneously withdrawing plunger to eliminate

as much headspace as possible before replacing the cap. The sample was submerged in the

reactor, which was pre-filled with silicon oil and held at 5 °C. The reactor was sealed with its

threaded cap before initiating pressurization. Once the set point for pressure was reached, water

flow was switched to the second, warmer water bath maintaining the preset incubation

temperature. Process time began once the reactor reached 95% of the temperature set point.

Figure 2.2 shows the pressure and temperature profile for a sample treated at 35 °C at 100 MPa

for 0 and 300 s. To determine 100% residual activity for a time of 0 s (t0), the reactor

immediately began cooling once 95% of the temperature set point was reached. Once the reactor

reached 15 °C depressurization was initiated as cooling continued to 5 °C. Samples were

immediately removed and analyzed for activity within two minutes of treatment.

24

Figure 2.2. Pressure (○) and temperature (□) profiles for a sample treated at 35 °C at 100 MPa

for 300 s and pressure (●) and temperature (■) profiles for a sample treated at 35 °C at 100 MPa

for 0 s

Processing Conditions

Samples were treated from 0.1 (control) to 150 MPa at 50 MPa increments at 25 °C, 35

°C, or 45 °C. Arrhenius equation was used to calculated the activation energy at each pressure.

At least four different treatment times were performed on samples to produce an approximate

80% reduction in residual activity after the longest processing time for each pressure-temperature

combination. Experiments were completed in a randomized block design, blocked by

temperature, and treatments completed in duplicate. Temperature was blocked due to the

impracticality of adjusting the water bath temperature between measurements, as doing so would

drastically increase overall experiment time.

Activity Measurements

0

20

40

60

80

100

120

140

0

10

20

30

40

50

60

70

80

0 100 200 300 400 500 600

Pre

ssure

(M

Pa)

Tem

per

ature

(ºC

)

Time (s)

25

A bi-enzymatic assay adapted to a 96-well microplate was used to measure the activity of

POX spectrophotometrically at 37 °C [61]. H2O2 produced by the POX-catalyzed reaction

(equation 2.2) was monitored at 550 nm by the POD-catalyzed conversion of EHSPT to

quinoneimine dye. (equation 2.3).

𝑃𝑦𝑟𝑢𝑣𝑎𝑡𝑒 + 𝑂2 + 𝑃𝑖𝑃𝑂𝑋→ 𝐴𝑐𝑒𝑡𝑦𝑙𝑝ℎ𝑜𝑠𝑝ℎ𝑎𝑡𝑒 + 𝐶𝑂2 + 𝐻2𝑂2 (2.2)

2𝐻2𝑂2 + 4 − 𝐴𝑚𝑖𝑛𝑜𝑎𝑛𝑡𝑖𝑝𝑦𝑟𝑖𝑛𝑒 + 𝐸𝐻𝑆𝑃𝑇𝑃𝑂𝐷→ 𝑄𝑢𝑖𝑛𝑜𝑛𝑒𝑖𝑚𝑖𝑛𝑒 𝑑𝑦𝑒 + 4𝐻2𝑂 (2.3)

A stock solution of approximately 8 units/mL POX was prepared and stored in an amber glass

bottle at 5 °C. Each day, a 0.8-unit/mL working solution was prepared by diluting 0.1 mL of

stock with 0.9 mL 50 mM potassium phosphate buffer, pH 5.7 (enzyme diluent) in a 8-mL amber

glass vial. New stock solution was prepared approximately every three weeks during

measurement, once the activity of the fresh enzyme began to decrease. A reaction cocktail was

prepared according to Table 2.1, adapted from Sigma [61]. Peroxidase, FAD, and TPP were

prepared fresh each day. All other solutions, including pyruvate and buffers were prepared

beforehand. When not in use, all reagents were held at 4 °C and stored on ice during

experiments.

Table 2.1. Reaction cocktail composition for pyruvate oxidase activity assay

Reagent Volume (mL)

150 mM Potassium phosphate buffer, pH 5.9 2.0

7.4 mM 4-Aminoantipyrine 0.4

0.3% (w/v) EHSPT 0.4

3 mM TPP 0.4

0.15 mM FAD 0.4

15 mM EDTA 0.4

150 mM MgSO4 0.4

50 units/mL POD 0.6

Total 5.0

26

Four microwels were used for each assay. First, 250 µL of reaction cocktail and 50 µL of

300 mM pyruvate solution were added to each of the wells. The pre-heated microplate mixed the

plate by swirling for 30 s, then monitored the absorbance for 5 min to ensure constant

absorbance before addition of enzyme. The plate was then ejected, 10 µL of enzyme diluent were

added to the first two wells to serve as a blank and 10 µL of enzyme sample were added to the

second two. The absorbance was then monitored for 20 min, with measurements occurring

approximately every 10 s, for a total of 121 measurements for each well. After each assay, the

data from the four cells was transferred into an Excel spreadsheet where the average of the two

blank wells was subtracted from the average of the two test wells to account for any non-

enzymatic oxidation of the substrate. Once this corrected absorbance was calculated for each

replication, the slope of absorbance versus time was calculated every 10 successive data points.

The activity (abs/time) was calculated as the maximum slope from each replicate. Residual

activity for a given replicate was calculated by comparing the activity of that replicate to the

average activity of the two replicates at time zero of the same treatment. The activity of fresh

enzyme was recorded daily to ensure no loss of activity of stock enzyme between comparable

treatments. All absorbance measurements were corrected for path length. Actual enzyme activity

(units/mL) was calculated according to equation 2.4.

𝑈𝑛𝑖𝑡𝑠 𝑚𝐿 𝐸𝑛𝑧𝑦𝑚𝑒⁄ = (∆𝐴550 𝑇𝑒𝑠𝑡−∆𝐴550 𝐵𝑙𝑎𝑛𝑘)×0.31

(36.88)(0.01)(2.4)

where 0.31 is the total volume in milliliters of the assay, 36.88 is the millimolar extinction

coefficient of quinonimine dye, 0.01 is the milliliter volume of enzyme used in the assay, and

one unit of POX is defined as producing 1.0 µmole of H2O2 per minute under assay conditions

(pH 5.7, 37 °C) [61].

27

Based on the good fit of the linear regression of natural logarithm of residual activity

versus time (figure 2.9), first-order kinetics were used to calculate the rate constant of

inactivation (k) at each selected temperature and pressure.

The Arrhenius equation (equation 2.5) was used to calculate the activation energy at each

pressure from the first-order rate constants of inactivation.

ln(𝑘) = (−𝐸𝑎𝑖

𝑅×1

𝑇) + ln (𝑘𝑇0) (2.5)

Where k is the rate constant, T is the absolute temperature, R is the ideal gas constant (8.314 J

mol-1 K-1), Eai is the activation energy, and 𝑘𝑇0 is the rate constant at a reference absolute

temperature. The “i” was added to the symbol Eai to denote that it is the activation energy of

inactivation as opposed to the activation energy of activation.

Activation volume of POX was calculated using Eyring’s equation (Equation 2.6) at each

temperature tested with the rates of inactivation:

ln(𝑘) = (∆𝑉≠

𝑅𝑇×𝑃) + ln (𝑘𝑃0) (2.6)

where k is the rate constant, ΔV≠ is the activation volume, P is pressure, 𝑘𝑃0is the rate constant at

reference pressure P0.

Standard error of the linear regression was reported for inactivation rates, activation energies,

and activation volumes.

Original Design with Plastic Pouches

The initial methods of this study were based on similar studies on the protective effects of

HHP on the stability of alcohol oxidase [60], xanthine oxidase, and glucose oxidase [58]. As

with enzyme samples in the aforementioned studies, POX samples were originally contained in

plastic pouches. These pouches were fabricated from low-density polyethylene zip-top bags,

using a heat sealer to divide the bags into 1.9 cm2 portions, which were cut into individual

28

squares so that the top edge of each square was left unsealed. A 100 µL aliquot of POX working

solution was added to each pouch slowly to allow for space between the bulk of the solution and

the edge of the bag, which would be heat sealed before the sample was added to the HHP system.

Initial results were inconsistent and poorly reproducible, with severe activity loss

occurring between fresh POX and unheated pressurized samples. Aside from pressure, heat from

the sealer and pouch material were considered as possible factors for loss in activity.

To account for potential loss from the heat sealer, pouch design and handling methods

were adjusted. The pouch design was modified to allow for more space between the heat sealer

and enzyme solution. In this new, narrower design, the pouches were cut into 1.3 x 3.8 cm

rectangles, with the open end on one of the narrower edges. Figure 2.3 shows a comparison of

the two pouch designs. A smaller volume (50 µL) of POX working solution was added to these

pouches to allow for a greater space between the heating element of the sealer and the enzyme

solution. Additionally, the pouches were kept on ice constantly from filling to addition to the HP

reactor, other than when they were being sealed at which point they were handled with tweezers

to avoid any temperature change from body heat. To examine the loss in activity due to heat

from the heat sealer, environment, and manual handling, the activity of POX sealed in the new

pouches and handled to minimize external heat was compared with fresh enzyme solution. These

samples were all pipetted into the pouches and sealed, then their activity was recorded.

29

Unsealed Edge

Figure 2.3. Comparison of original (left) and newer (right) pouch design to scale

After only modest improvements in enzyme activity retention from adjustments to pouch

design and handling, the possible effect of sample container material was considered. Enzyme

solution was added to an amber glass vial, one of the plastic pouches, and a high-density

polypropylene centrifuge tube, all stored in a covered ice bucket. The plastic pouch was left

unsealed so that any inactivating effects would be due to container material and not incidental

heating. The activity of the POX solution was recorded from each container at 0, 25, and 50 min.

Unlike the original activity assay protocol that involved two test wells and two blank wells, each

sample was added to a single well, along with a single blank well so that all samples could be

measured at the same time.

Unsealed Edge

Sample

(100 µL)

Sample

(50 µL)

30

Pouch Study Results

Residual activity measurements using the original pouch showed highly inconsistent

results. Table 2.2 details residual activity for POX treated with 100 MPa without raising

temperature using the square pouches. On average, virtually no activity was lost over the short

treatment time of 75 s, however, the standard deviation (n = 3) between the triplicates was

extremely high. Relative to the fresh POX working solution, which remained in an amber glass

vial, the loss for POX treated in the original pouches was severe (75 – 80%).

Table 2.2. Residual activity for pyruvate oxidase treated using original pouch design at 100 MPa

at 5 °C relative to both 0 s treatment time and fresh working enzyme solution (n=3)

Residual Activity (%)

Time (s) Relative to 0 s Relative to Fresh

0 100 ± 44 % 21 ± 9 %

75 116 ± 68 % 25 ± 14 %

Based on the results from experiments using the original pouch design, it was considered

that enzyme activity was being affected by sources of heat outside of the reactor, namely heat

from the sealer, but possibly also atmospheric and body temperature. However, new pouch

design and handling methods resulted only in slight improvements in activity loss. Table 2.3

outlines residual activity values for these unpressurized samples relative to fresh POX solution.

31

Table 2.3. Residual pyruvate oxidase activity for samples sealed in new pouches relative to fresh

enzyme activity over time

Sample 1 Sample 2 Sample 3

Residual Activity (%)

Time (min)

61%

0

49%

30

12%

60

At best, there was still at least a 60% loss in activity using the new pouch design and handling

technique. It is important to note that all three samples were prepared at the same time, and each

remained on ice while the previous sample’s activity was recorded, meaning there was

approximately 30 – 35 min between activity assays. Results indicated that POX loses activity

over time as it remained in the pouch, as the activity of Sample 1 was recorded immediately after

being sealed in the pouch, Sample 2 remained in the pouch for approximately 30 min, and

Sample 3 for at least 60 min.

The comparison of activity over time for POX contained in a plastic pouch, glass vial,

and hard plastic tube (Figure 2.4) revealed that, over the span of 50 min, the activity of POX was

reduced by more than half when stored in the plastic pouches, while the samples stored in glass

or hard plastic decreased from 88% to 70% and 79% to 61% respectively. Data for the POX

stored in the vial at 25 min was excluded due to pipetting error into the single microwell. This

experiment was not replicated.

32

Figure 2.4. Residual activity relative to fresh enzyme over time for pyruvate oxidase in low-

density polyethylene pouch ( ), high density polypropylene tube ( ), and amber glass vial ( )

In a study that compared the retention of eight radiolabeled proteins in containers made

of different plastic and glass materials commonly found in lab settings, Goebel-Stengel, et al.

[62] concluded that selecting the proper can improve protein retention dramatically. In their

comparison of polypropylene and polystyrene, the two most commonly-used plastics in

laboratories, retention for polypropylene was as good or better than polystyrene. In the case of

cholecystokinin, the difference in retention was 86 vs 16% for the two plastics respectively.

Specific protein-plastic interactions are difficult to predict due to the variability in peptide

structure between different proteins. Based on data from these preliminary studies, all future

experiments in this study were completed using high-density polypropylene syringes.

Native and Inactivated Enzyme Comparison

Native polyacrylamide gel electrophoresis (PAGE) was performed to attempt to

determine whether the tetrameric structure of pyruvate oxidase broke into subunits (dimers or

monomers) once completely inactivated with pressure. The gels were submerged in a 25-mM

Tris, 192 mM glycine running buffer. To prepare the inactivated sample, 100 µL of POX stock

0%

20%

40%

60%

80%

100%

120%

0 25 50

Res

idual

Act

ivit

y R

elat

ive

to F

resh

Enzy

me

Time (min)

33

solution (6 units mL-1), prepared in 50 mM, pH 5.7 potassium phosphate buffer, was added to a

syringe and treated at 45 °C at 150 MPa for 30 min. This treatment was determined based on

prior data to achieve complete inactivation of the enzyme. Before loading onto the gels, 50 µL of

fresh enzyme stock solution, inactivated solution, and potassium phosphate buffer were each

separately mixed with an equal volume of native running buffer. For each sample, 40 µL of

solution, containing approximately 4 µg of protein, was added to two wells. A 200-V potential

was applied to the system for 60 min. Afterwards, the gel was removed and stained with

Coomassie blue for 30 min on a shaker platform, then rinsed with water for 30 min on the same

platform.

To further understand the effect of the inactivation treatment on the protein, the

absorbance spectrum between 240 – 350 nm was recorded using a NanoDrop™ One

Microvolume UV-Vis spectrophotometer for both inactivated and native enzyme samples, as

well as a sample of 50 mM phosphate buffer treated at 150 MPa at 45 °C for 30 min to detect

any potential contaminants from the plastic syringe that may absorb within the spectrum. For

each of the three samples, 2 µL was pipetted directly onto the loading stage and the optical piece

was lowered such that the sample droplet rested underneath.

34

Results and Discussion

Activity Assay

Preliminary activity measurements revealed an initial lag phase lasting around 10 min

before any change in absorbance was observed. While not reported in POX from Aerococcus sp.,

POX from E. coli also experiences an initial lag phase [63]. Shortly after the lag phase,

absorbance increased linearly for all samples. When compared to fresh enzyme samples, there

was a decrease in activity for samples treated for 0 s. Figure 2.5 compares the absorbance over

time for POX treated 35 °C at 0.1 MPa for 0 and 600 s, as well as fresh POX working solution

from the same day. The slope (absorbance/time) decreases as treatment time increases from 0 to

600 s, and a slight decrease in slope occurs between fresh POX sample and the sample treated for

0 s. This would indicate not only a loss in POX activity over time, but also some loss occurring

after the initial heating/pressurization associated with the t0 treatment.

Figure 2.5. Corrected absorbance measurements for individual replicates of fresh pyruvate

oxidase (black) compared with samples treated at 0.1 MPa at 35 °C for 0 (dark grey) or 600 s

(light grey).

0

0.05

0.1

0.15

0.2

0 5 10 15 20

Abso

rban

ce

Time (min)

35

The loss in activity due to the t0 treatment at each pressure and temperature combination relative

to fresh enzyme stock solution is shown in Figure 2.6. The process of bringing samples to the

temperature and pressure setpoint reduced activity relative to fresh in all cases. From 0.1 to 100

MPa, there was a consistent reduction in activity of about 20% at all temperatures, with an

increase in reduction at 150 MPa. Higher reduction in enzyme activity between fresh enzyme

and those treated for 0 s occurred at lower temperatures at 150 MPa.

Figure 2.6. Residual activity for t0 treatments relative to fresh enzyme at 25 ( ), 35 ( ), or

45 °C ( )

Additionally, there is a delay between the heating of the reaction vessel and the interior of

the syringes. By comparing the two temperatures (Figure 2.7), it is evident that the syringes take

approximately an additional two minutes to reach the temperature setpoint after the reactor.

0%

20%

40%

60%

80%

100%

120%

0 50 100 150

Res

idual

Act

ivit

y R

elat

ive

to

Fre

sh (

%)

Pressure (MPa)

36

Figure 2.7. Reactor (open symbol) versus syringe (closed symbol) temperature at 25 (light grey),

35 (dark grey), or 45 °C (black)

Effect of HHP on Stability

Pyruvate oxidase residual activity decreased over time at all pressure-temperature

combinations. Figure 2.8 shows the loss in residual activity for samples treated at 0.1, 50, 100,

and 150 MPa at all three temperatures. Residual activity generally decreased as pressure

increased.

0

10

20

30

40

50

0 50 100 150 200 250 300

Tem

per

ature

(°C

)

Time (s)

37

Figure 2.8. Residual activity for POX at 25 (A), 35 (B), or 45 °C (C) at 0.1 ( ), 50 ( ), 100 ( )

or 150 ( ) MPa

First order models were used to describe the relationship between residual activity of

POX and treatment time. First order kinetics are, to our knowledge, the most commonly used

model for the rate of thermal and pressure inactivation of enzymes, as in the case of glucose

oxidase [58], pectin methylesterase [64], and alcohol oxidase [60]. Such models provided a good

-30%

20%

70%

120%

0 1000 2000 3000 4000 5000 6000 7000 8000

Res

idual

Act

ivit

y (

%)

Time (s)

0.0%

50.0%

100.0%

150.0%

0 500 1000 1500 2000 2500 3000 3500 4000 4500 5000 5500

Res

idual

Act

ivit

y (

%)

Time (s)

0.0%

50.0%

100.0%

150.0%

0 500 1000 1500 2000

Res

idual

Act

ivit

y (

%)

Time (s)

A

B

C

38

fit for residual activity as a function of treatment time and allowed for meaningful comparison to

other data from the literature

First order rate constants of inactivation (kinact) are reported in Table 2.4. At 25 °C, the

rate of inactivation increased with pressure at pressures above 50 MPa, with there being no

statistically significant change between atmospheric pressure and 50 MPa. The rate of

inactivation for 50 MPa at 35 °C was significantly smaller than 0.1 and 100 MPa, while there

was no significant difference between 0.1 and 100 MPa. The poor coefficient of determination of

determination (R2) for the 50 MPa treatment at 35 °C can be partially explained by the very

small slope for the rate of inactivation, as seen in Figure 2.9.

39

Table 2.4. First order rate of pyruvate oxidase inactivation ± standard error determined by the

linear regression of residual activity versus treatment time.

Pressure Temperature (°C)

25 R2 35 R2 45 R2

(MPa) kinact (min-1)

0.1 0.003a ± 0.001 0.42 0.004a ± 0.003 0.22 0.089a ± 0.007 0.95

50 0.004a ± 0.002 0.50 0.001* ± 0.001 0.08 0.065a ± 0.021 0.49

100 0.027 ± 0.002 0.95 0.006a* ± 0.002 0.57 0.138 ± 0.013 0.90

150 0.182 ± 0.035 0.72 0.177 ± 0.019 0.90 0.433 ± 0.045 0.91

a Represents statistically similar treatments as determined by ± standard error calculated from the

linear regression for each temperature.

* Indicates treatments completed with a newer batch of enzyme

At 45 °C, the rate of inactivation generally increased with pressure, except for a small

decrease from 0.1 to 50 MPa, but this decrease was not statistically significant. There was a large

increase in rate of inactivation between 100 and 150 MPa at all temperatures. While pressure-

induced inactivation of POX from Aeroccocus has not been explored in the current literature,

fluorescence data shows that in POX from E. coli, the cofactors FAD and TPP dissociate from

the enzyme before unfolding occurs as pressure approaches 220 MPa [13].

40

Figure 2.9. Logarithm of residual activity at 25 (A), 35 (B), or 45 °C (C) at 0.1 ( ), 50 ( ),

100 ( ) or 150 ( ) MPa

To investigate a possible discrepancy between enzyme batches, a series of experiments

using the new batch were completed comparing the residual activity overtime between 0.1 and

50 MPa at all three temperatures. The results from these supplemental experiments (Table 2.5)

show a similar response between enzyme batches. The residual activity values for the new batch

1.00

2.00

3.00

4.00

5.00

0 1000 2000 3000 4000 5000 6000 7000 8000ln(R

esid

ual

Act

ivit

y,

%)

Time (s)

1.000

2.000

3.000

4.000

5.000

0 1000 2000 3000 4000 5000 6000ln(R

esid

ual

Act

ivit

y,

%)

Time (s)

1.000

2.000

3.000

4.000

5.000

0 200 400 600 800 1000 1200 1400 1600ln(R

esid

ual

Act

ivit

y,

%)

Time (s)

A

B

C

41

was with those calculated based on kinact values for the old data at the same treatment times. The

difference between batches was mostly on the same order as the variability between replicates in

the same batch, but variability increased at 45 °C. The pressure/temperature treatment profiles

for these supplemental experiments (Figures A.1 – A.10) show that the treatments are both

consistent and reproducible.

42

Table 2.5. Residual activity data for confirmation experiments using new pyruvate oxidase batch

compared to residual activity for the old batch calculated based on kinact.

Treatment Time (s) Rep 1 Rep 2 Average

Old Batch

(Calculated)

Difference

(New Batch -

Old)

0.1 MPa 25 °C

0 98.6% 101.4% 100.0%

3000 94.2% 88.9% 91.5% 94.9% -3.4%

50 MPa 25 °C

0 98.9% 101.1% 100.0%

3000 100.2% 86.6% 93.4% 92.4% 1.0%

0.1 MPa 35 °C

0 93.4% 106.6% 100.0%

3000 102.8% 107.7% 105.3% 92.2% 13.1%

50 MPa 35 °C

0 94.2% 105.8% 100.0%

3000 98.0% 124.6% 111.3% Same Batch

0.1 MPa 45 °C

0 103.9% 96.1% 100.0%

1000 56.8% 34.2% 45.5% 70.0% -24.5%

50 MPa 45 °C

0 103.5% 96.5% 100.0%

1000 87.9% 113.3% 100.6% 77.0% 23.6%

Based on the data for this batch-to-batch comparison, there was insufficient evidence to discard

residual activity values based on new batch POX from future calculations.

Activation Energy of Inactivation

The apparent activation energy of inactivation (Eai) was calculated using the Arrhenius

equation. Table 2.6 reports Eai for 0.1 to 150 MPa at increments of 50 MPa. With every increase

in pressure of 50 MPa, the apparent Eai decreased by slightly less than a half, meaning that the

enzyme’s sensitivity to temperature increases with pressure over the observed range of pressure.

43

Table 2.6. Apparent activation energy of inactivation ± standard error determined by the linear

regression of ln(k) versus 1/T

Pressure Eai R2

(MPa) (kJ mol-1)

0.1 135 ± 66 0.81

50 100 ± 15 0.33

100 62 ± 113 0.24

150 33 ± 21 0.71

Activation energy of inactivation for POX decreased by around 75% as pressure was

increased from 0.1 to 150 MPa. The decrease in Eai for POX was more dramatic over a smaller

pressure range than many other enzymes from similar studies. Comparatively, Eai for glucose

oxidase dropped from 378.1 to 281.0 kJ/mol between 0.1 – 300 MPa [58] and xanthine oxidase

saw a decrease from 181.7 to 97.0 kJ/mol over 0.1 – 300 MPa [59]. In alcohol oxidase, Eai for

the thermoresistant fraction of the enzyme saw no significant change from 105.1 to 101.3 MPa

over a pressure range of 40 – 200 MPa. Even for some enzymes inactivacted by pressure, unlike

those previously mentioned, the degree of change in Eai was often less than that observed for

POX. The activation energy for pectinmethylesterase from navel oranges decreased from 177 to

95 kJ/mol over range of 100-750 MPa [64] while Eai for plum polyphenoloxidase just halved

from 130.6 to 62.7 over a dramatic pressure increase from 0.1 to 900 MPa [65]. Compared to the

plum polyphenol oxidase and pectinmethyesterase from navel orange, which saw their Eai halved

over a pressure range of 650 and 900 MPa respectively, the Eai of POX was reduced by 75% over

a range of only 150 MPa. This rate in reduction of Eai in POX as pressure increases is nearly an

order of magnitude greater, demonstrating its remarkable sensitivity to thermal inactivation as

pressure increases.

44

Activation Volume

Eyring’s equation was used to calculate activation volume (ΔV≠). Table 2.7 reports

activation volume for 25, 35, or 45 °C. For 25 and 35 °C, ΔV≠ was very similar. However, at 45

°C the activation volume fell by over 50% relative to the other two treatments, suggesting that

pyruvate oxidase is far more sensitive to pressure at 45 °C than the two lower temperatures.

Different trends in ΔV≠ could be observed in previously-studied oxidases. For xanthine oxidase,

ΔV≠ roughly tripled from 9.5 to 28.9 cm3/mol as temperature increased from 55.0- 70 °C [59]. In

glucose oxidase, ΔV≠ changed from 22.8 to 45.8 cm3/mol over a temperature range of 58.8 – 80.0

°C, with the greatest ΔV≠ of 57.0 cm3/mol occurring at 74.5 °C [58]. Additionally, polyphenol

oxidase from strawberry (Fragraria ananassa) saw an increase in ΔV≠ from -53.41 to -2.81

cm3/mol between 45 and 65 °C [66]. As with each of these enzymes, an increase in ΔV≠ is

associated with higher stability, indicating that more pressure is needed to achieve inactivation as

stability increases. In the case of POX, the decrease in ΔV≠ at 45 °C from 25 and 35 °C is within

reason, given the much higher rates of inactivation at 45 °C than those at the two lower

temperatures.

45

Table 2.7 . Activation volume ± standard error determined by the linear regression for each

temperature

Temperature ∆V≠ R2

(°C) (cm3 mol-1)

25 71 ± 12 0.95

35 68 ± 43 0.56

45 29 ± 13 0.72

* Indicates values calculated excluding new batch data, for which standard error and R2 could not

be calculated

When comparing the rate constant of inactivation versus pressure at each temperature (Figure

2.10) the plots form a gentle curve at the bottom of which sit 50 MPa. Based on the projected

shape of these plots, it is unlikely that a significantly lower inactivation rate could be achieved

just above or below 50 MPa.

Figure 2.10. Rate constant of inactivation versus pressure at 25 ( ), 35 ( ), and 45 °C ( )

0.000E+00

1.000E-03

2.000E-03

3.000E-03

4.000E-03

5.000E-03

6.000E-03

7.000E-03

8.000E-03

0 50 100 150 200

k (

s-1)

Pressure (MPa)

46

Native and Inactivated Enzyme Comparison

The stained polyacrylamide gel (Figure 2.11) showed that both the native and a portion of

the inactivated enzyme traveled the same distance, signifying that they had the same mass/charge

ratio. The bands for the inactivated enzyme were much fainter than those for the native, and there

was protein that remained in the top of the wells for the inactivated samples. This suggests that a

portion of the protein may have formed an aggregate too large to enter the gel, while the

remaining portion has returned to a native conformation upon depressurization. In a study on

another homotetrameric enzyme, urate oxidase, Girard, et al. [55] observed the enzyme structure

under high pressure and measured activity before and after pressurization. They observed that

above 150 MPa urate oxidase approaches a threshold of irreversible damage to the quaternary

structure, wherein a portion of the enzyme aggregates while the rest of the still-soluble enzyme

remains in a native, tetrameric state. Evidence of unfolding in pyruvate oxidase can be seen

spectrophotometrically (Figure 2.12). The “peak broadening” as absorbance increases, especially

below 270 nm, could be a result of cofactors being released as the enzyme unfolds. Thiamine

pyrophosphate exhibits a strong peak around 240-250 nm at neutral pH [67]. The release of

buried TPP from POX could explain the increase in absorbance on the lower end of the UV

spectra. The lack of absorbance in the blank sample would imply no contamination from the

syringe during processing.

47

Figure 2.11. Relative distance traveled through 4-20 % polyacrylamide gel by native and

inactivated pyruvate oxidase samples.

Figure 2.12. Ultraviolet spectra for Native (black), Inactivated (grey), and blank (dotted) samples

-0.2

0

0.2

0.4

0.6

0.8

1

1.2

220 240 260 280 300 320 340 360

Abso

rban

ce

Wavelength (nm)

Native Inactivated Blank

48

Conclusion

In conclusion, both HHP and temperature largely had an additive inactivating effect on

pyruvate oxidase from Aerococcus species, with pressure offering little to no protection against

thermal inactivation except for a small protective effect at 50 MPa at 35°C. Given the findings of

this study and the current literature on POX stability, HHP is not a viable technique for

stabilizing POX from Aerococcus sp. for the fabrication of biosensors.

49

Chapter Three

Final Comments

Overview

Data showed that material from the low-density polyethylene containers diminished

enzyme activity. This could either be through POX adsorbing to the plastic surface or due to the

plastic having an inactivating effect on the enzyme. Either way, these results further highlighting

the general importance of selecting appropriate containers when working with enzymes. While

the high-density polypropylene syringes did not have the same inactivating effect of POX, there

was a delay between the reactor temperature that was recorded and that of the temperature of the

samples.

Future Work

Further work is required to determine if lower pressures (around 50 MPa) can be utilized

in conjunction with other techniques to increase the feasibility of biosensors based on pyruvate

oxidase. Results from this study show some potential for pressures beneath what is usually

considered high pressure (100-700 MPa), perhaps in combination with other stabilization

techniques. Given the slight stabilization seen at 50 MPa in both the original and new batch data,

it would be useful to examine the effect of pressure between 15 – 75 MPa at a similar

temperature range to this study.

Based on data from native PAGE and UV-spectroscopy, it may be possible that POX,

like urate oxidase, maintains its native conformation until approaches a threshold of irreversible

50

aggregation. More sophisticated methods in which pyruvate oxidase is directly observed under

high pressure are needed to confirm the accuracy of this model, and to confirm if the internal

cavities play a role in the process of unfolding and aggregation. This could be achieved using

protein simulations informed by x-ray crystallography and/or fluorescence spectroscopy under

high pressure.

51

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57

APPENDIX

SUPPLEMENTAL EXPERIMENT TEMPERATURE AND PRESSURE PROFILES

The following figures compare the temperature and pressure profiles of samples from the

supplemental experiments using the new batch of POX, to show consistency in treatment

between replicates. Profiles for samples treated a 0.1 MPa at 25 °C for 0s have been omitted, as

the combination of low pressure, short time, and low pressure erroneously trigger the LabView

program to cease recording as treatment continues. The file for the second replication treated at

50 MPa at 45 °C for 1000 s was mistakenly overwritten.

Figure A.1. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 0.1 MPa at 25 °C for 3000 s

58

Figure A.2. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 50 MPa at 25 °C for 0 s

Figure A.3. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 50 MPa at 25 °C for 3000 s

Figure A.4. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 0.1 MPa at 35 °C for 0 s

59

Figure A.5. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 0.1 MPa at 35 °C for 3000 s

Figure A.6. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 50 MPa at 35 °C for 0 s

Figure A.7. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 50 MPa at 35 °C for 3000 s

60

Figure A.8. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 0.1 MPa at 45 °C for 0 s

Figure A.9. Pressure (closed symbol) and temperature (open symbol) for samples one (dark grey)

and two (light gray) for supplemental sample treated at 0.1 MPa at 45 °C for 1000 s

Figure A.10. Pressure (closed symbol) and temperature (open symbol) for samples one (dark

grey) and two (light gray) for supplemental sample treated at 50 MPa at 45 °C for 0 s