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The Pharmaceutical stability and formulation of plasmid DNA

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Title:
The Pharmaceutical stability and formulation of plasmid DNA
Creator:
Poxon, Scott William, 1971-
Publication Date:
Language:
English
Physical Description:
xii, 116 leaves : ill. ; 29 cm.

Subjects

Subjects / Keywords:
DNA ( jstor )
DNA damage ( jstor )
Endotoxins ( jstor )
Foams ( jstor )
Freeze drying ( jstor )
Gene therapy ( jstor )
Lipids ( jstor )
Liposomes ( jstor )
pH ( jstor )
Plasmids ( jstor )
DNA -- chemistry ( mesh )
Department of Pharmaceutics thesis Ph.D ( mesh )
Dissertations, Academic -- College of Pharmacy -- Department of Pharmaceutics -- UF ( mesh )
Drug Stability ( mesh )
Endotoxins ( mesh )
Plasmids -- chemistry ( mesh )
Research ( mesh )
Transfection ( mesh )
Genre:
bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Thesis:
Thesis (Ph.D.)--University of Florida, 1999.
Bibliography:
Bibliography: leaves 107-114.
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Scott William Poxon.

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University of Florida
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University of Florida
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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
Resource Identifier:
030253967 ( ALEPH )
51638680 ( OCLC )

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THE PHARMACEUTICAL STABILITY AND FORMULATION OF PLASMID DNA









By



SCOTT WILLIAM POXON


















A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY


UNIVERSITY OF FLORIDA



1999








































This dissertation is dedicated to my wife,

Stephanie Poxon, without whose understanding and

patience I could not have accomplished this task. As

well, I would like to thank my parents and the other

students in the Hughes laboratory for their help and

support.



















ACKNOWLEDGMENTS

I would like to acknowledge my committee members,

Dr. Jeffrey Hughes, Dr. Gayle Brazeau, Dr. Edwin

Meyer, Dr. Michael Schwartz, and Dr. Ian Tebbett for

their critical reading of this manuscript, support and

contribution to this project. The Parenteral Drug

Association Foundation Biotechnology Grant, the

National Institutes of Health (NIH P01-AG 10485), and

the University of Florida Gene Therapy Center also

financially supported these studies.






















iii
















TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ........................................... iii

LIST OF FIGURES ................................... ...... vi

ABSTRACT ................................................. xi

1 INTRODUCTION ............................................ 1

Specific Aims and Hypotheses............................ 1
Background and Significance............................. 2
Chemical and Physical Stability of DNA.................. 5
Background ........................................... 5
Glycosidic Bond ...................................... 7
Phosphodiester Bond ......... ......... ........ ...... 9
Steric Effects .......................... ............. 10
Oxidative Damage ....................................... 10
Lyophilization of DNA................................. 11
Endotoxin Contamination of DNA......................... 13
Conclusions ............................................15

2 PHYSICAL AND CHEMICAL STABILITY OF PLASMID DNA ......... 16

Background ...... .......................... ........... 16
Materials and Methods..................................21
Plasmid Purification ............................... 21
Liposome Preparation ............................... 22
In vitro Coupled Transcription-Translation ..........23
Transfection Efficiency Assay ......................23
DMED Assay .......................................... 25
H Solution Stability Study ......................... 25
Aemperature Stability Study ........................26
Statistical Analysis ................................ 27
Results and Discussion................................. 27

3 FORMULATION OF PLASMID DNA: THE EFFECT OF
LYOPHILIZATION ON PLASMID DNA STABILITY................ 34

Introduction. .............................. ............ 34

iv









Materials and Methods.................................. 38
Plasmid Purification ................................ 38
Liposome Preparation .............................. 39
Transfection Efficiency Assay ...................... 40
Differential Scanning Calorimetry Analysis .......... 41
DMED Assay ..................... ... ......... ............... 42
Analysis of Hyperchromic Effect ............. ....... 42
Oxidative Analysis ...................... .. ....... 43
Circular Dichroism Analysis ........................ 44
Statistical Analysis ................................ 44
Results and Discussion................................. 44
Conclusion............................................. 61

4 CHARACTERIZATION OF ENDOTOXIN AND CATIONIC LIPOSOME
INTERACTION ........................................... 64

Introduction ........................................... 64
Materials and Methods.................................. 67
Plasmid Purification ............................... 67
Liposome Preparation ................................ 68
Transfection Efficiency Assay ...................... 68
Anisotropy Assay ............................... .. .. 70
Endotoxin Dephosphorylation ........................ 71
Endotoxin Assay .......................... .. ......... 71
Cell Viability Assay ................................ 72
Statistical Analysis ................................ 73
Results................................................ 73
Discussion ............................................. 81

5 FOAM FRACTIONATION AS A METHOD TO SEPARATE ENDOTOXIN
FROM RECOMBINANT BIOTECHNOLOGY PRODUCTS................ 88

Introduction............................................. 88
Methods ................................................ 91
Plasmid Purification ............................. ... 91
Foam Fractionation .................................. 92
Plasmid and FITC-Endotoxin Gel Analysis .............92
Surface Activity Determination ...................... 93
Particle Size Analysis ............................. 93
Endotoxin Assay ...................... .... .......... 93
Protein Analysis ..................................... 94
Statistical Analysis ........................... ....... 94
Results and Discussion...... .............. ............ 94
Conclusions ................................ ............. 103

LIST OF REFERENCES ...................................... 107

BIOGRAPHICAL SKETCH .................................... 115

v











LIST OF FIGURES


Figure page

1. Major sites for chemical degradation of DNA.
Filled arrows point to potential oxidative damage
sites, while open arrows point to potential
hydrolytic damage sites. ..............................6

2. Glycosidic cleavage of cytosine to an aldehyde
under alkaline or acidic conditions followed by
beta elimination to cleave the phosphate
backbone. .......................... ................... 7

3. Comparison of cotransfection (A) and single
plasmid (B) concentration vs activity. Mean +
SEM, n=4. .............................................28

4. Effect of incubation at elevated temperature for
3 weeks on the functional activity of plasmid DNA
as measured via coupled transcription-
translation. RLU+SEM, n=3, 370C is significantly
different than 75 and 950C (p<0.01) via Fisher's
PLSD. ................................................. 29

5. Effect of incubation at various pHs for 3 weeks
on the functional activity of plasmid DNA as
measured via coupled transcription-translation.
RLU+SEM, n=6, *=p<0.05 via Fisher's PLSD
compared to pH 7 ......................................30

6. Effect of incubation in pH 3, 2.5 mM citrate
buffer on plasmid DNA in a 0.8% agarose gel,
stained with ethidium bromide. .......................31

7. Effect of citrate buffer concentration on pRL-
CMV plasmid degradation rate at 500C. Mean +
SEM, n=5, r2>0.98 for all fit lines. ..................32

8. Effect of lyophilization (FD) on plasmid DNA
activity, with various amounts lyoprotectant to
DNA (w/w). Average + SEM. n=5 for all
treatments, = FD DNA significantly lower than
Control DNA. p<0.05 via Schefe's multiple
comparison T-test. ...................... ............46


vi









9. Representative thermal analysis scan of salmon
sperm DNA ............................................ 49

10. Effect of lyoprotectants on the melting
temperature of salmon-sperm DNA. Mean + SEM,
n=3, = significantly different than unprotected
control DNA (p < 0.05) via Fisher's PLSD. 50

11. Effect of lyophilization and subsequent
rehydration on the melting of plasmid DNA. Mean +
SEM, n=3 ..............................................51

12. Lyophilized plasmid DNA samples, rehydrated in
DI water and run an agarose gel. Samples:
marker, HindII digested lambda phage marker; FD,
lyophilized DNA without DMED treatment; FD DMED,
lyophilized DNA with DMED treatment; pH 3, DNA
incubated in pH 3.0, 2.5 mM citrate buffer for 15
minutes at 500C with no DMED treatment; pH 3
DMED, DNA incubated in pH 3.0, 2.5 mM citrate
buffer for 15 minutes at 500C with DMED treatment
before running on gel .................................52

13. Effect of lyophilization on plasmid form. Mean +
SEM, n=3. p > 0.05 via ANOVA ........................53

14. Effect of lyophilization on plasmid DNA
absorbance at 260 nm. Average + SEM. n=4, FD
DNA significantly higher than lyoprotected DNA
p<0.05 via ANOVA ..................................... 54

15. Wavelength circular dichroism scan of
lyophilized plasmid DNA (FD) compared to A-form
and B-form plasmid DNA. Each scan is the average
of three separate baseline subtracted scans. ...........55

16. Effect of lyoprotection on plasmid DNA
conformational change. Panels each signify a
separate protectant; A: lactose, B: glucosamine,
C: glucose-l-phosphate, and D: glucose. For each
panel Protectant:DNA 1, 5, 10, 20 and 40 or 1,
2, and 8 w/w ratios. Cntrl S "protectant" is
protectant:DNA solution (40:1 w/w) non
lyophilized. Cntrl FD and Cntrl S "without
protectant" are no protectant, lyophilzed or non
lyophilized respectively. n=10, mean + SEM. ......... 56



vii









17. Kinetics of plasmid conformational change after
rehydration. mean + SEM, n=3.. ..................... 58

18. Effect of rehydration time on plasmid DNA
activity. Mean + SEM. n=6. No significant
difference between 0 and 24 hours by T-test. ......... 59

19. HPLC analysis of hydrolyzed pRL-CMV plasmid DNA.
Panel A is a UV analysis. Peaks by retention
time: 2.76, solvent front, 3.79 cytosine, 7.85
guanine, 11.15 thymine, 13.00 unidentified, 17.89
adenine. Panel B is an ECD analysis. Peaks by
retention time 2,67 3.21 solvent front, 3.53 5-
OH cytosine, 3.82 cytosine, 4.78 plasticizer,
7.35 guanine, 7.92 8-OH guanine, 16.99 8-OH
adenine and adenine. ...................................60

20: Effect of lyophilization of plasmid DNA on
oxidative damage. Positive control was Fe+3
catalyzed oxidation. Mean + SEM, n=5, except
control n=3. = p < 0.05 via Fisher's PLSD
versus control. ........................................ 61

21. Effect of tube type on plasmid DNA bioactivity
after lyophilization. Cntrl = non-lyophilized
DNA, PP = polypropylene, SP = siliconized
polypropylene, PE = polyethylene. Mean + SEM,
n=5, = p < 0.05 via Fisher's PLSD versus
control, X = p < 0.05 versus PP and SP. ..............62

22. Enzyme activity corrected for total cellular
protein after transfection of luciferase plasmid
(lpg) in the presence of endotoxin, with and
without DOTAP : DOPE cationic liposomes. 0 with
lipid (2 ug/ml), U without lipid. RLU+SEM, n=4,
p<0.05 via one way ANOVA for effect of endotoxin
in the presence of cationic lipid. ................... 74

23. Effect of increasing DOTAP : DOPE liposomes
concentration with FITC-endotoxin held constant
(1000 EU) on anisotropy (r). 0 DI Water, m 0.5 M
NaC1, A 1.0 M NaC1, X 2.0 M NaCI. Inset: Free
FITC (22 ng) 1.0 M NaC1. Change in r+SEM, n=3,
p<0.05 using two way ANOVA for increase in r with
increasing lipid under all conditions. ............... 76




viii









24. Effect of increasing luciferase plasmid
concentration on anisotropy (r) with constant
FITC-endotoxin (1000 EU) and DOTAP : DOPE
liposomes (5 gg). Change in r+SEM, n=3, p<0.05
using two way ANOVA for decrease in r with
increasing plasmid. ..................................77

25. Effect of increasing lipid or dendrimer
concentration with FITC-endotoxin held constant
(1000 EU) on anisotropy (r). U Dendrimer
(Generation 2), 0 Dendrimer (Generation 4), X
lecithin liposomes (0.2 pm), A lecithin liposomes
(0.8 gm), Change in r+SEM, n=3, p<0.05 using two
way ANOVA for increase in r with increasing
concentration under all conditions. ..................78

26. Effect of increasing DOTAP : DOPE liposome
concentration with FITC-Endotoxin held constant
(10 gg) .............................................. 78

27. Effect of increasing incubation time of alkaline
phosphatase and unlabeled endotoxin (33 EU) on
anisotropy (r) with constant and NBD labeled
DOTAP : DOPE liposome (10 Vg). Change in r + SEM,
n=4, p>0.05 via one way ANOVA for decrease in
anisotropy over time ................................ 80

28. Effect of increasing lipid concentration with
endotoxin held constant (0.6 EU) on endotoxin
activity, EU/ml + SEM, n=3, p>0.05 via one way
ANOVA for effect of cationic lipid on endotoxin
activity. ............................................. 81

29. Effect of increasing cationic lipid (panel A)
and endotoxin (panel B) on COS-1 cell survival
via MTT assay. mean + SEM, n=5, p < 0.05 via
ANOVA for DOTAP : DOPE, p > 0.05 for endotoxin
via ANOVA. ................. ......................... 82

30. Effect of foam fractionation on FITC labeled
endotoxin levels (initial 100 pg/ml n=3) and BSA
(initial 910 gg/ml, n=3) in a plasmid DNA
solution, (25 ig/ml), mean + SEM. p>0.05 using
one way ANOVA for differences in endotoxin
concentration over time. p<0.05 for differences
in BSA concentration over time. .....................96


ix









31. Effect of foam fractionation on endotoxin
activity. Mean + SEM, p > 0.05 via ANOVA ............97

32. Effect of foam fractionation on physical
stability of plasmid DNA, molecular weight marker
X phage Hind III digest in lane 1; lanes 2 8,
fractionation time points 1 gg pDNA per lane, in
a 0.8% agarose gel stained with ethidium bromide. ....98

33. Surface tension (dynes/cm) of water, endotoxin,
FITC-endotoxin, and BSA (all 1 mg/ml), n=4, mean
+ SEM, = p<0.05 using Fisher's PLSD versus
water .................................................99

34. Volume weighted particle size of FITC-endotoxin
(1 mg/ml) at increasing HC1 concentration. Data
are shown as mean+SEM., p<0.05 via ANOVA, *=
p<0.05 using Fisher's PLSD verus 1 pM HC1 ........... 100

35. Surface tension (dynes/cm) of FITC-endotoxin (1
mg/ml) at various HC1 concentrations, n=4, mean +
SEM, = p<0.05 using Fisher's PLSD versus 0 pm
HC1 .................................................. 101




























x
















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of
the Requirements for the Degree of Doctor of Philosophy


THE PHARMACEUTICAL STABILITY AND FORMULATION OF PLASMID DNA


By



Scott W. Poxon



May, 1999



Chairman: Jeffrey Hughes, Ph.D.
Major Department: Pharmaceutics


This research examined the stability and formulation

of plasmid DNA from a pharmaceutical perspective. Plasmid

DNA has the potential to be used as a drug, which could

replace missing or damaged proteins that cause disease. As

well, plasmid DNA could be used as the basis for new types

of vaccines. However, the potential for this new drug can

not be realized without research into the formulation and

stability of plasmid DNA. These studies examined the



xi









stability of plasmid DNA with respect to environmental

factors such as temperature and pH, showing that plasmid

DNA is exceedingly stable under many conditions.

Furthermore, this research compared the stability of

plasmid DNA in both solution and solid lyophilized states.

These studies demonstrated that the lyophilization process

damages DNA through what appears to be a conformationally

induced denaturation, although the lyophilized plasmid DNA

is more stable at elevated temperature than plasmid DNA in

solution. Finally, it was ascertained that a potential

contaminant of plasmid DNA, endotoxin, can decrease DNA

transfection efficiency through a competitive electrostatic

interaction with a common DNA delivery vector, cationic

liposomes. It was also established that the endotoxin

contaminant can not be removed from plasmid DNA using foam

fractionation. However, the foam fraction method can

successfully be used to separate endotoxin from amphiphilic

protein products. This research should help to support the

future pharmaceutical development of plasmid DNA as a

therapeutic modality.










xii















CHAPTER 1
INTRODUCTION


Specific Aims and Hypotheses

The overall goal of this project was to determine the

impact of environmental factors on plasmid DNA stability.

Using structural and biofunctional assays, the stability

and potential mechanisms of plasmid DNA instability were

investigated. This information was then assessed with

respect to the pharmaceutical development of plasmid DNA in

a therapeutic modality.

Four specific hypotheses were investigated by this

project. The first hypothesis was that biofunctional assays

are more sensitive than structural assays, since damage at

one base in the encoding region would not be easily

detected by conventional structural methods, but could

result in a functionally inactive product. Secondly, it

was hypothesized that environmental factors will affect the

stability of plasmid in solution. The third hypothesis was

that the stability of lyophilized plasmid DNA to

environmental factors will be higher compared to plasmid

DNA in solution. However, the lyophilization process may


1






2


damage plasmid DNA via conformational strain caused by the

removal of the DNA hydration sphere. Finally, this project

tested the hypothesis that increasing levels of endotoxin

contamination would result in reduced functionality of

plasmid DNA in tissue culture models.

To test these hypotheses, the following were the

specific aims of this work:

1. To establish and validate a sensitive and specific assay

to quantitate the structural and biofunctional stability

of plasmid DNA.

2. To determine the optimum storage conditions for plasmid

DNA in solution.

3. To determine the influence of the hydration sphere on

optimum storage conditions for lyophilized plasmid DNA.

4. To determine the effect of plasmid DNA endotoxin

contamination on functionality using specific assays.


Background and Significance

DNA, or deoxyribonucleic acid, is the genetic material

of life. It is used to pass information from one

generation to the next, supplying the set of genes needed

for the manufacture of further structures in the organism.

DNA functions via its capacity to encode a large variety of

proteins, where specific sequences of nucleotides in DNA






3


encode different proteins. Structurally, DNA consists of

two antiparallel strands whose sequences are made up of

chemically linked subunits, consisting of a nitrogenous

base (purine or pyrimidine) attached to a pentose sugar

linked together by a phosphate backbone. Each nucleic acid

contains one of four types of nitrogenous base: two

purines, adenine and guanine, and two pyrimidines, cytosine

and thymine. In mammals, chromosomes make up the discrete

unit of the genome. Each chromosome consists of a long

duplex DNA complexed with protein.

Another form of DNA, plasmid DNA, is an autonomous

unit that can exist inside a bacterial cell's cytoplasm.

These extrachromosomal, double stranded, circular molecules

of DNA are self-replicating. During the 1970s, many

artificial plasmids were constructed in the laboratory

utilizing fragments from naturally occurring plasmids, and

are commonly used as vectors for recombinant DNA work and

gene therapy.

Since DNA is the genetic material, mutations to DNA

can potentially have serious consequences. A change in the

sequence of DNA causes an alteration of the coded protein,

which may cause mutational inactivation. Mutations may

either be inherited or induced. In either case,

replacement of the mutated DNA via gene therapy can result





4


in the production of competent protein. This makes gene

therapy one of the most exciting and rapidly advancing

areas of medicine. Great strides have been recently made

in the development of DNA as a therapeutic agent including

the successful injection and expression of plasmid DNA into

animals (Nabel et al. 1989; Nabel et al. 1990; Nabel et al.

1992; Stewart et al. 1992). These initial successes have

quickly led to human clinical trials (Nabel et al. 1993;

Caplen et al. 1994).

Although there has been some initial investigation

into the potential production methods for pharmaceutical-

grade DNA (Horn et al. 1995; Durland, and Eastman 1998),

little information is available concerning plasmid DNA

stability in a pharmaceutically acceptable dosage form.

For instance, shelf life has not been determined in any

potential dosage form, and it is commonly assumed that DNA

is stable as either an ethanol precipitate or a frozen

buffered solution at neutral to slightly basic pH (Maniatis

et al. 1982). However, neither postulate has been

thoroughly investigated with respect to the pharmaceutical

development of plasmid DNA.






5


Chemical and Physical Stability of DNA


Background

The reactivity of the phosphate backbone, the

deoxyribose, and the nitrogenous bases that make up the

individual components of DNA has been well documented

(Shabarova, and Bogdanov 1994). Initial research in this

area determined how bases may react with exogenous

electrophilic agents, resulting in heterocyclic

halogenation or nitration. Other electrophilic reactions

include methylation and oxidation. Reactions with amine

containing nucleophilic reagents may also result in

substitutions under basic conditions (Shabarova, and

Bogdanov 1994). Of these reactions, oxidation is most

likely of concern in long-term DNA storage since the

plasmid isolation process should not result in the addition

of exogenous electrophiles (Finnegan et al. 1996).

However, plasmid isolation may result in the addition of

activated oxygen species through air saturated buffers or

the use of organic solvents for extraction resulting in

subsequent oxidative damage (Finnegan et al. 1996).

Although these previous reactions have generally been

characterized experimentally using nucleic acid monomers,

it is possible that the reactivity of plasmid DNA may





6

increase or decrease significantly from the monomers due to

steric factors associated with the primary and secondary

structures of plasmid DNA. Several specific areas of the

DNA have been extensively studied for their lability under

physiologic conditions. The sites susceptible to

hydrolytic attack and oxidative damage in plasmid DNA can

be seen in Figure 1.





Cytosine Guanin /
SH2 "........' NA -O-

5"' "'NI N INN'N

-o0 ......H2N 9

"........ H2 PN
9 5
-0- N 0 Adenine
0- Thymine



3'

Figure 1: Major sites for chemical degradation of DNA.
Filled arrows point to potential oxidative damage sites,
while open arrows point to potential hydrolytic damage
sites.






7


Glycosidic Bond

The glycosidic bond is highly stable under neutral or

basic conditions, but is extremely sensitive to acid

hydrolysis. Acid hydrolysis of the glycosidic bond on

nucleotides occurs rapidly with purines (k=10-4sec-1) and

more slowly with pyrimidines (k=10-8sec-1) at pH 1.0, 370C,

since the purines are better leaving groups than

pyrimidines (Lindahl 1993). Under physiologic conditions,

it has been predicted that mammalian cells undergo 2,000 to

10,000 cases of hydrolytic depurination followed by repair

every day (Lindahl 1993). In pharmaceutical storage of

plasmid DNA, however, no repair mechanisms exist, bringing

concern about potential degradation of the glycosidic bond.

NH12
5' 5' 5'

ON O --- HOH 0 ----0 O


O-6 H H -O





Figure 2: Glycosidic cleavage of cytosine to an aldehyde
under alkaline or acidic conditions followed by beta
elimination to cleave the phosphate backbone.

After depurination by cleavage of the glycosidic bond,

an aldehyde forms at the C1 of deoxyribose, which can then






8


undergo beta elimination as shown in Figure 2 (McHugh, and

Knowland 1995). This aldehyde form exists in equilibrium

with the cyclic depurinated hemiacetal form, with about 1%

of the base-free sugar residue in the aldehyde form at any

one time. Even without chemical catalysis, the weakened

DNA chain would undergo the elimination process within a

few days, rendering the stored plasmid DNA non-functional

by blocking DNA polymerase.

It has been shown that deamination of the bases may

also occur in alkaline conditions, resulting in potential

loss of the encoded protein's functionality. This

deamination process occurs mainly via acid-catalysis under

physiologic conditions (Lindahl 1993). This, however, is a

slow reaction, with the half-life of an individual cytosine

residue in single stranded DNA extrapolated to 200 years

(Frederico et al. 1990). The hydrolytic deamination

reaction slows further with double stranded DNA to a half-

life of about 30,000 years for each cytosine residue

(Frederico et al. 1990). Deamination occurs at a slower

rate than depurination, and is therefore less likely to be

the rate-determining step in plasmid degradation during

storage.






9


Phosphodiester Bond

During prolonged exposure to high temperatures, DNA

will progressively melt via first order kinetics followed

by heat-induced hydrolysis of the phosphodiester bond with

an expected 3,000 fold increase in DNA decay at 1000C as

compared to 370C (Lindahl 1993). At temperatures above

100C the DNA is very unstable because of both its chemical

nature to hydrolyze and the problem of retaining the

hydrogen bonding between the two DNA strands at high

temperatures. It has also been suggested that increased

pressure could help stabilize DNA, since the melting

temperature of the double helix is 100C higher at 5,000

atmospheres than at 1 atmosphere (Lindahl 1993).

The phosphodiester bond is another potential site for

damage in plasmid DNA. It may be broken by beta

elimination as well as by acid hydrolysis at a pH below 3.

The phosphodiester backbone can also be cleaved by

nucleases and by oxidative degradation, but is otherwise as

stable at neutral and basic pH as the glycosidic bond and

the amine groups on the bases. This explains the reasoning

behind current DNA storage paradigms of freezing DNA at

neutral to slightly basic pH.






10


Steric Effects

Positive supercoiling can act as a barrier to

deamination, depurination and subsequent beta elimination.

Protection via supercoiling is due to a change in the

plasmid's tertiary structure that results in torsional

twisting of the DNA structure. This twisting causes the

duplex to cross itself in space and is responsible for the

increased pH needed to denature circular DNA as compared to

linear DNA. Supercoiling can also protect against thermal

DNA degradation (Marguet, and Forterre 1994), suggesting

that condensation of plasmid DNA may increase stability

during storage.


Oxidative Damage

Oxidative damage of plasmid DNA may be a problem

inherent in long-term storage, particularly if the DNA was

exposed to hydroxyl or superoxide radicals (Lindahl 1993).

Exposure to activated oxygen radical is ubiquitous inside

cells, where DNA repair enzymes exist that reduce the

damage to cellular DNA. In long-term storage of plasmid

DNA, these repair enzymes will not exist, so oxidative

damage can accrue resulting in non-functional protein

product. Damage can potentially occur during plasmid DNA

isolation with solvents such as phenol and chloroform, and






11


air-saturated buffers (Lindahl 1993; Finnegan et al. 1996).

Exposure to activated oxygen can cause the formation of

formamidopyrimidines, purines with opened imidazole rings,

that are non-coding residues. As well, hydroxyl radicals

can react with guanine, forming 8-hydroxyguanine, which

base pairs with adenine rather than cytosine. Therefore,

transversion mutations will be generated after replication.

Another common oxidative-induced DNA damage is the

generation of ring saturated pyrimidines. In this case,

losing the 5,6 double bond causes the loss of planar ring

structure leaving a non-coding base residue (Lindahl 1993).


Lyophilization of DNA

Lyophilization of plasmid DNA may be a preferred form

of storage, potentially imparting a longer shelf life and

greater stability against heat-induced degradation. /

Experimental findings support this by suggesting that

biological macromolecules demonstrate an increase in

stability from the frozen to the lyophilized state (Crowe

et al. 1990). The exact mechanism underlying this

difference has not been fully elucidated, but most likely

relates to the nonfreezable water associated with most

biomolecules, including nucleic acids. It is well known

that the structure and conformational states of DNA are






12


critically dependent on hydration level (Umrania et al.

1995; La Vere et al. 1996). Several forms of DNA secondary

structure are known and, under normal physiological

conditions, DNA is found in the B-form. This form has a

major and a minor groove, with 10 base pairs per turn. The

A-form is more compact, with 11 base pairs per turn, and is

seen when the presence of 2'hydroxyl groups on the ribose

prevents formation of the B-form during complexation with

RNA, and under conditions of high salt concentration, or

low humidity (<75%), as expected under lyophilized

conditions. While the A and B-forms are both right handed

helixes, the Z-form is a left handed helix and has the most

base pairs per turn of any of the forms, and only one

groove. This form has been seen under high salt

concentrations and probably does not exist in vivo but

could potentially be seen in low concentrations under the

high salt conditions seen with lyophilization. The

secondary structure of the DNA may affect stability and

functionality.

Lyophilization causes the removal of the hydration

sphere around a molecule. For DNA, it appears that there

are approximately 20 water molecules per nucleotide pair

bound most tightly to DNA that do not form an ice-like

structure upon low temperature cooling. Upon DNA






13


dehydration at 0% relative humidity only five or six water

molecules remain (Falk et al. 1963; Lith et al. 1986; Tao,

and Lindsay 1989). Lyophilization may increase the

stability of DNA under long-term storage, but may also

cause some damage upon the initial lyophilization process,

potentially through changes in the DNA secondary structure.

Agents that can substitute for nonfreezable water, such as

trehalose, can demonstrate cryoprotective properties for

DNA and other molecules during lyophilization of intact

bacteria (Rudolph et al. 1986; Israeli et al. 1993). Other

cryoprotective agents, such as polyols, amino acids, sugars

and lyotropic salts are preferentially excluded from

contact with protein surfaces but are also capable of

stabilizing enzymes during lyophilization by undefined

mechanisms (Carpenter, and Crowe 1989). It is possible

that agents that act as cryoprotectants to proteins may

also act to stabilize nucleic acids.


Endotoxin Contamination of DNA

Since plasmid DNA is typically produced by bacteria,

endogenous bacterial products can potentially contaminate

the final plasmid DNA product. Typical problem compounds

in plasmid DNA preparations include endotoxin, a cell wall

component that is pyrogenic in man, and DNase, a ubiquitous






14


bacterial enzyme that can degrade the final plasmid

product. Endotoxin from the cell wall of Gram-negative

bacteria, such as the E. coli typically used to prepare

plasmid DNA, may be the most serious concern for gene

therapy because of its extreme toxicity. Several plasmid

DNA preparation methods have been examined for level of

endotoxin contamination (Weber et al. 1995).

Endotoxin has a relatively low lethal dose for 50% of

tested animals (LDso) in rats of 3 mg per kg (Shibayama et

al. 1991) and an LD50 in dogs of only 1 mg per kg (Fletcher,

and Ramwell 1980). The endotoxin, or lipopolysaccharide, of

E. coli consists of a polysaccharide component and a

covalently bound lipid component termed lipid A. The lipid

A portion on the endotoxin is biologically active and can

cause a number of pathophysiological effects including

fever, hypotension, intravascular coagulation and death

(Fletcher, and Ramwell 1980; Aida, and Pabst 1990;

Rietschel et al. 1993; Xing et al. 1994). The endotoxins

are acute inflammatory mediators, elevating levels of

cytokines (Xing et al. 1994). Furthermore, an increasing

level of endotoxin contamination has been shown to decrease

transfection efficiency using DOTAP : DOPE (dioleoyl

glycero trimethylammonium propane : dioleoyl glycero

phosphoethanolamine) cationic liposomes to deliver plasmid





15


DNA (Weber et al. 1995). Besides having an easy way to

detect endotoxin, there are several ways to remove

endotoxin from DNA samples. Phase separation using Triton

X-114 has been used to reduce endotoxin levels for both

protein and plasmid DNA (Aida, and Pabst 1990; Manthorpe et

al. 1993). The use of these methods to reduce endotoxin

contamination may be of importance in the potential

usefulness of plasmid DNA.


Conclusions

It is important to understand the effects of the

formulation parameters such as pH, temperature, and buffer

composition on plasmid DNA stability. Furthermore, the

differences between solution and lyophilized plasmid

stability must be taken into account to determine the

optimal plasmid DNA storage conditions. This research will

address the stability of plasmid DNA by examining plasmid

structural integrity and biofunctionality. This

distinction is important, since current structural methods

will not always detect changes in the functionality of the

final protein product that can be produced by damage to the

plasmid DNA. Using the information provided by these

studies, it should be possible to develop a rational basis

for the plasmid DNA manufacturing process.















CHAPTER 2
PHYSICAL AND CHEMICAL STABILITY OF PLASMID DNA


Background

Before the benefits of gene therapy can be realized on

a large scale, the pharmaceutical community needs to

ascertain the optimal storage conditions for plasmid DNA so

as to prolong shelf life to the greatest possible extent.

While a great deal of work has gone into the study of

individual nucleotides, little has been accomplished to

date regarding the bioactivity of plasmid DNA, or the

ability of plasmid DNA to code for a functional protein, as

a marker for shelf life. Other methods have been used to

examine DNA stability, including agarose gel

electrophoresis, HPLC, Southern blotting, and PCR (Niven,

Pearlman et. al 1998; Strege and Lagu 1991; Thierry,

Lunardi-Iskandar et. al 1995). However, a bioactivity based

method will detect changes not only in physical stability

of plasmid DNA but also will detect cellular changes in the

DNA handling, including, but not limited to, transport,

transcription, and translation.




16






17


The stability of the subunits of plasmid DNA, the

individual purine and pyrimidine bases, and their

attachment to the ribose sugar and phosphate backbone are

fairly well understood. The glycosidic bond is highly

stable under neutral or basic conditions, but is extremely

sensitive to acid hydrolysis. Acid hydrolysis of the

glycosidic bond on nucleotides occurs rapidly with purines

(k=10-4sec-) and more slowly with pyrimidines (k=10-ssec-1)

at pH 1.0, 37C, since the purines are better leaving

groups than pyrimidines (Lindahl 1993). Under physiologic

conditions, it has been predicted that mammalian cells

undergo 2,000 to 10,000 cases of hydrolytic depurination

followed by repair every day (Lindahl 1993). In

pharmaceutical storage of plasmid DNA, however, no repair

mechanisms exist, bringing concern about potential

degradation of the glycosidic bond.

After depurination by cleavage of the glycosidic bond,

an aldehyde forms at the Cl of deoxyribose, which can then

undergo beta elimination as shown in Figure 2 on page 7

(McHugh, and Knowland 1995). This aldehyde form exists in

equilibrium with the cyclic depurinated form, with about 1%

of the base-free sugar residue in the aldehyde form at any

one time. Even without chemical catalysis, the weakened

DNA chain will undergo the elimination process within a few






18


days. This would render the stored plasmid DNA non-

functional by blocking DNA polymerase.

It has been shown that deamination of the bases may

occur in alkaline conditions, resulting in potential loss

of the encoded protein's functionality. This process can

also occur via acid hydrolysis under physiologic

conditions, where the half-life of an individual cytosine

residue in single stranded DNA has been extrapolated to 200

years (Frederico et al. 1990). This hydrolytic deamination

reaction slows further with double stranded DNA to a half-

life of about 30,000 years for each cytosine residue

(Frederico et al. 1990). This reaction occurs at a slower

rate than depurination and is therefore less likely to be

the rate-determining step in plasmid degradation during

storage. Depurination rates may be effected by changes in

the formulation of plasmid DNA.

During prolonged exposure to high temperatures, DNA

will progressively melt via first-order kinetics followed

by heat-induced hydrolysis of the phosphodiester bond, with

an expected 3,000 fold increase in DNA decay at 1000C as

compared with 370C (Lindahl 1993) This rate increase

essentially follows the 100C rule with a 3-fold rate

increase per 100C temperature increase. At temperatures

above 1000C the DNA is very unstable because of both its





19


chemical nature to hydrolyze and the problem of retaining

the hydrogen bonding between the two DNA strands at high

temperatures. It has also been suggested that increased

pressure could help stabilize the DNA, since the melting

temperature of the double helix is 100C higher at 5,000

atmospheres than at 1 atmosphere (Lindahl 1993).

The phosphodiester bond is another potential damage

site for plasmid DNA. It may be broken by beta elimination

as well as by acid hydrolysis at a pH below 3. The

phosphodiester backbone can also be cleaved by nucleases

and by oxidative degradation, but is otherwise as stable at

neutral and basic pH as the glycosidic bond and the amine

groups on the bases. This explains the reasoning behind

current DNA storage paradigm of frozen DNA at neutral to

slightly basic pH.

Positive supercoiling can act as a barrier to

deamination, depurination and subsequent beta elimination.

This supercoiling protection is due to a change in the

plasmid's tertiary structure that results in torsional

twisting of the DNA structure. This twisting causes the

duplex to cross itself in space and this conformationally

induced steric protection is responsible for the increased

pH needed to denature circular DNA as compared to linear

DNA. Supercoiling can also protect against thermal DNA






20


degradation (Marguet, and Forterre 1994). If supercoiling

exerts a protective effect, then it follows that DNA

delivery systems that further condense supercoiled plasmid

DNA may additionally increase stability during storage. As

well, supercoiled plasmid DNA may consequently be more than

the monomers that were originally used to determine DNA

stability.

While it is possible to examine plasmid DNA stability

through examination of any of the above individual

degradation pathways, a better way to examine DNA stability

may be through the examination of plasmid DNA bioactivity,

by assaying for enzymatic activity of the plasmid's encoded

gene. Assaying for activity of the plasmid DNA will allow

a comprehensive determination of degradation that will

include all types of damage that affect the expressed gene

rather than just the several specific types of damage

previously mentioned. The activity of the plasmid can be

determined through one of several bioactivity assays.

First, plasmid DNA can be assayed using an in vitro coupled

transcription-translation system. The in vitro system

allows a one tube determination of activity that is not

dependent on any particular type of cell system for

expression. Rather, the in vitro system includes all the

necessary enzymes and cofactors to transcribe the plasmid






21


DNA to mRNA and to further translate the mRNA to protein,

which can then be assayed. This method's main disadvantage

is due to variability induced by the 'stop-time" of the

reaction. If the reaction is stopped at different times,

the resulting enzymatic activity will be effected.

The second method for determining the bioactivity of

the plasmid is using a mammalian cell based tissue culture

transfection assay. This method requires that the plasmid

DNA be complexed with a delivery agent before it is

transducted into a mammalian cell line. The cell line will

then express the protein of interest, which can then be

assayed. Several main types of DNA damage are expected, as

discussed previously. This study will examine the

stability of plasmid DNA using both types of activity

assays.


Materials and Methods


Plasmid Purification

Vectors pGL3 plasmid, pSPluc+, and pRL-CMV (Promega,

Madison, WI), respectively encoding for photinus

luciferase, photinus luciferase and renilla luciferase,

were grown in E. coli DH5a cells (Promega, Madison, WI).

The transformed bacteria were cultivated in Lura Bertina

(LB) broth containing 100 ug/ml ampicillin to select for 3-






22


lactamase encoding plasmid. The plasmids were isolated via

an alkaline lysis method and purified using a silica slurry

column (Wizard Plus Megapreps, Promega, Madison, WI). The

plasmid was stored in TE buffer (10 mM Tris-HCl, 1 mM EDTA,

pH 7.4).


Liposome Preparation

Lipids were obtained from Avanti Polar Lipids

(Alabaster, AL). Cationic DOTAP : DOPE (dioleoyl glycero

trimethylammonium propane : dioleoyl glycero

phosphoethanolamine) liposomes were prepared using the

hand-shaking method (New 1990). DOTAP: DOPE (10 mg DOTAP,

10 mg DOPE) was added to 10 ml of chloroform and introduced

into a 250 ml round bottom flask. The chloroform was

evaporated using a rotary evaporator at 600C. The lipid

film was hydrated with 10 ml of distilled water and shaken

for 30 minutes at 600C. Lipids were sized by extrusion, six

times through 600 nm polycarbonate filters (Poretics Corp,

Livermore, CA). Sizes were confirmed with a laser light

scattering particle sizer using volume-weighted

distribution (Nicomp 380ZLS, Santa Barbara, CA).





23


In vitro Coupled Transcription-Translation

The coupled transcription-translation assay was

carried out using a commercially available kit (Promega,

Madison, WI). Essentially, pGL3 plasmid DNA (100 pg) was

incubated for 60 minutes at 300C with a proprietary mixture

containing wheat germ extract, RNA polymerase, amino acids,

and a ribonuclease inhibitor. The resulting protein

product, luciferase, was then assayed for functionality

using a luminescence spectrophotometer (Monolight 2010,

Analytical Luminescence Laboratory, San Diego, CA). The

light output was measured for 10 seconds and the results

integrated to yield the activity.


Transfection Efficiency Assay

Untreated pGL3 (0.05 pg/ml) and treated pRL-CMV (0.5

pg/ml) were combined with cationic lipid mixture (DOTAP :

DOPE 1:1, 2 pg/ml each) and incubated for 15 minutes to

allow interaction of the anionic plasmid with the cationic

liposomes. The 10:1 concentration ratio was as suggested

by the manufacturer to achieve a linear relationship

between treated renilla luciferase plasmid concentration

and renilla luciferase / photinus luciferase activity ratio

by decreasing crosstalk between the promoters. These

mixtures were then co-transfected into COS-1 cells (ATCC,






24


Rockville, MD) that had been plated 24 hours earlier at

3x104 cells/ml in 24 well tissue-culture dishes. All

transfections were effected using serum free DMEM

(Dulbecco's Modified Eagle's Medium) supplemented with

penicillin G (100 units/ml) and streptomycin (100 pg/ml).

The cells were subsequently incubated for 5 hours in a

humidified, 5% CO2 incubator at 37C. The media was

aspirated and the cells washed with PBS (phosphate buffered

saline) before being replaced by DMEM supplemented with 10%,

FBS (Fetal Bovine Serum), penicillin G (100 units/ml) and

streptomycin (100 pg/ml). The cells were incubated for an

additional 19 hours, washed with 1 ml PBS, and then lysed

with Passive Lysis buffer (200 pL, Promega, Madison, WI).

Photinus luciferase and renilla luciferase activities

were determined by analyzing 5 pL of the lysate using the

Dual Luciferase Assay kit (Promega, Madison, WI) and a

luminescence spectrophotometer (Monolight 2010, Analytical

Luminescence Laboratory, San Diego, CA). The light output

was measured for 10 seconds and the results integrated to

yield the activity. All pRL-CMV, renilla luciferase

plasmid, measurements were standardized to the ratio of

renilla luciferase (rluc treated) activity to photinus

luciferase (luc control) activity. Essentially, the ratio






25


of rluc/luc is linear to the concentration of rluc added to

COS-1 cells. Therefore, a linear regression of rluc/luc

ratio at various concentrations of rluc and inactivated

rluc (cut with restriction enzymes), with luc held

constant, allows a determination of the apparent

concentration of the treated pRL-CMV plasmid.


DMED Assay

Treated pGL3 DNA was incubated with N,N'-

dimethylethylenediamine (100 mM, pH 7.4, 30 minutes at

37C), cleaving abasic sites via 3-elimination (McHugh, and

Knowland 1995). The resultant DNA was then visualized

using a 0.8% agarose gel. The gel was electrophoresed at 1

V/cm for 16 hours to affect good separation between the

supercoiled, relaxed circular and linear forms of the

plasmid DNA. The gel was then stained with ethidium

bromide and photographed on an UV light box using a DC40

digital camera (Kodak, Rochester, NY).


pH Solution Stability Study

The effect of pH upon plasmid DNA bioactivity was

studied using a multiple component buffer (Schrier et al.

1993). Poly-B buffer, equimolar sodium citrate, sodium

succinate, Tris, HEPES, imidazole, histidine, and glycine

at various pHs, was added to pSP6-luc plasmid DNA to a






26


final concentration of 10 mM each salt and 0.25 mg/ml

plasmid DNA. This mixture was incubated at room

temperature for three weeks, with the assumption that pH

did not change over time. The plasmid was then

precipitated using 70% ethanol, and the pellet rehydrated

in TE buffer (10 mM Tris-HC1, 1 mM EDTA, pH 7.4) before

analysis by the coupled transcription-translation method.

The potential for buffer catalysis of plasmid DNA was

examined using a citrate buffer at four different

concentrations, all at pH 3, 500C. Plasmid, pRL-CMV, was

incubated with the various buffers, with aliquots withdrawn

over a course of five hours. The aliquots were returned to

neutral pH by the addition of excess TE buffer, pH 7.4.

The pH was verified to be greater than 7.0 using litmus

paper. These samples then were assayed using the

cotransfection method.


Temperature Stability Study

The effect of temperature upon plasmid DNA bioactivity

was examined after a three-week incubation at four

temperatures: 25, 37, 75 and 950C. Plasmid samples, pSP6-

luc, were stored in the humidified incubator at 1 mg/ml in

TE buffer in sealed 1 ml cryogenic screw-top vials. After






27


the three-week period, all samples were cooled and assayed

using the coupled transcription-translation method.


Statistical Analysis

Statistical analysis between the various treatments

was conducted using analysis of variance and Fisher's

protected least significant difference (PLSD) post-hoc T-

tests where appropriate (StatView v4.5, Abacus Concepts,

Berkley, CA), with p<0.05 considered statistically

significant.


Results and Discussion

The luciferase cotransfection method used two plasmids

coding for two different enzymes (fixed concentration

photinus and varying concentration renilla luciferase) to

help control for variability by comparing the ratio of the

two enzymes activity after transfection. The raw data was

analyzed by comparing only the activity of the renilla

luciferase. The luciferase cotransfection method of

normalizing transfection efficiency was compared to single

transfection, raw, non-normalized data shown in Figure 3.

It can be seen that the normalized cotransfection data

shows a better fit and shows less variability than the

single plasmid raw data. The percent coefficient of

variation (CV) and percent bias seen with both methods at






28


each measured concentration can be seen in Table 1. The

trend toward lower CV at plasmid concentrations above 0.13

gg / ml with the cotranfection suggests that the luciferase

cotransfection method is more sensitive than the single

plasmid method. Another trend is systematic bias below the

fit line, suggesting that the normalized method is more

reproducible than the single plasmid luciferase methods.

A 5 B o _
R2 0.9982 R' 0.9575
4 48



I 02--


0.0 0.1 0.2 0.3 0.4 0.5 0.0 0.1 0.2 0.3 0.4 0.5
pRL-CMV (ug) pRL-CMV (ug)

Figure 3: Comparison of cotransfection (A) and single
plasmid (B) concentration vs activity. Mean + SEM, n=4.


Table 1: Comparison of variance and bias between the
luciferase cotransfection and single plasmid methods


% CV % Bias
jg DNA Cotransfection Single Cotransfection Single
0.03 42 27 21 8
0.06 57 40 20 25
0.13 21 29 4 26
0.25 21 20 3 27
0.50 4 38 0 9


Extended incubation of pSP6-luc plasmid DNA for three

weeks at elevated temperature can result in a significant

decrease in the bioactivity of the treated plasmid at





29


elevated temperatures (Figure 4). This degradation would

be expected through heat-induced hydrolysis of the

phosphodiester bond. The bioactivity assay demonstrated a

three order of magnitude decrease in plasmid DNA activity

after incubation at 950C, which is in line with an expected

3,000 fold increase in DNA decay at 1000C compared to 370C

as previously reported in the literature (Lindahl 1993).



106


105


j 104


103


10


0
25 37 75 95
Storage Temperature (C)
Figure 4: Effect of incubation at elevated temperature for
3 weeks on the functional activity of plasmid DNA as
measured via coupled transcription-translation. RLU+SEM,
n=3, 370C is significantly different than 75 and 95C
(p<0.05) via Fisher's PLSD.

The effect of pH on pSP6-luc plasmid DNA bioactivity

was also examined (Figure 5). It can be seen that the





30


plasmid DNA remained exceedingly stable across a wide pH

range. Degradation was seen only at pH 1 and 2. This is

not suprising, since the glycosidic bond is reported in the

literature to be highly stable under neutral or basic

conditions, but is reported to be extremely sensitive to

acid hydrolysis (Lindahl 1993). Furthermore, the

phosphodiester bond may be broken by beta elimination as

well as by acid hydrolysis at a pH below 3. The loss of

bioactivity seen at low pH is most likely due to a

combination of the two degradation pathways.



106

105


104

n, 103

102

10



0
1 2 4 6 7 8 10
Storage pH
Figure 5: Effect of incubation at various pHs for 3 weeks
on the functional activity of plasmid DNA as measured via
coupled transcription-translation. RLU+SEM, n=6, *=p<0.05
via Fisher's PLSD compared to pH 7





31


Using the DMED assay to examine the mechanism of

acidic plasmid DNA degradation, it can be seen that nearly

100% of the plasmid DNA is depurinated at one or more sites

per plasmid molecule after 15 minutes (Figure 6). This

degradation rate is reasonable compared to published rates

with a T1/2 of 16 minutes at pH 1.0, 370C for depurination

(Lindahl 1993).


Standard DMED




OC
NL



SC


X 0 15 30 60 120 240 X 0 15 30 60 120 240

Minutes Minutes

Figure 6: Effect of incubation in pH 3, 2.5 mM citrate
buffer on plasmid DNA in a 0.8% agarose gel, stained with
ethidium bromide.

It was also interesting to note that besides

undergoing a pH dependent degradation, plasmid DNA may also

be subject to buffer catalysis. Statistically different

degradation rates were seen when incubating plasmid DNA,






32


pRL-CMV, in a 500C, pH 3.0 solution, buffered with various

amounts of citrate (Figure 7). Degradation rates appear to

be first-order exponential and increase with citrate buffer

concentration through an undetermined mechanism.


3

2.5

2 ~*.5 mM
S0 01 mmM
C 1.5 A A2mM

1 \

0.5 1..
0 5 10 15 20 25 30
Incubation Time (min.)


Figure 7: Effect of citrate buffer concentration on pRL-CMV
plasmid degradation rate at 500C. Mean + SEM, n=5, r2>0.98
for all fit lines.

In conclusion, plasmid DNA is exceedingly stable in

solution over a wide range of pH and temperature extremes.

However, the choice of buffer is of more importance than

has typically been considered in the past. Based on the

literature review and the data presented here, it is most

advisable to formulate plasmid DNA at neutral to slightly

basic conditions, using previously tested buffers. With

buffers that exhibit protective effects, a high

concentration of buffer would be preferable. With buffers

that that exhibit buffer catalysis, a low buffer






33


concentration that will minimize any potential buffer

effects should be used.















CHAPTER 3
FORMULATION OF PLASMID DNA: THE EFFECT OF LYOPHILIZATION ON
PLASMID DNA STABILITY


Introduction

Gene therapy is one of the fastest growing areas in

therapeutics. While rapid progress has occurred in this

field, the pharmaceutical community has not addressed all

concerns in detail. Of particular interest is the final

dosage form. It is well understood that naked DNA

introduced into a patient's circulatory system does not

reach enough of the appropriate cells and therefore has

little chance of affecting most disease processes

(Friedmann 1997). This has led to the development of a

number of gene delivery vectors, such as cationic

liposomes. Liposomes are considered one of the more

promising systems for use in gene delivery (Xu, and Szoka

1996). Unfortunately, cationic liposomes complexed with

plasmid DNA are not stable for long-term storage,

undergoing aggregation over time (Anchordoquy et al. 1997).

There have been several studies with non-ionic

liposomes, which may apply to the stability of the



34





35


DNA/cationic liposome mixture during the lyophilization

process. Lyophilization of non-ionic liposomes in the

presence of carbohydrates has been cited as one of the most

promising ways to keep the liposome stable under long term

storage (Williams, and Polli 1984). The lyophilization

process, without lyoprotectants, can lead to the leakage of

the inner aqueous phase due to liposomal fusion and phase

separation of liposomal membranes during drying and

rehydration (Crowe et al. 1988). Using carbohydrates as a

lyoprotectant will prevent mechanical rupture of the

liposomal membrane, caused by ice crystals, during the

freezing process and will prevent membrane disruption

during drying and rehydration by maintaining the membrane

in a flexible state (Ozaki, and Hayashi 1997).

Although DNA is complexed with cationic liposomes, not

entrapped, it has been shown that the liposome/DNA complex

requires lyoprotectants to maintain transfection efficiency

after lyophilization (Anchordoquy et al. 1997). Moreover,

other gene delivery systems containing polyethylenimine,

polylysine, and adenovirus particles require cryoprotection

to maintain transfection efficiency (Talsma et al. 1997).

However, it is unclear if this effect is due only to

stabilization of the liposomal component of the mixture or

if the plasmid DNA component may itself be damaged by






36


lyophilization in the absence of carbohydrate

lyoprotectants.

Lyophilized plasmid DNA may be a preferred form of

storage, potentially imparting a longer shelf life and

greater stability against heat-induced degradation, since

biological macromolecules demonstrate an increase in

stability from the frozen to the lyophilized state (Crowe

et al. 1990). The exact mechanism underlying this

difference has not been fully elucidated, but most likely

relates to the nonfreezable water associated with

biomolecules, including nucleic acids. It is well known

that the structure and conformational states of DNA are

critically dependent on hydration level (Umrania et al.

1995; La Vere et al. 1996).

Several forms of DNA secondary structure are known and

under normal physiological conditions most DNA is found in

the B-form. The B-form has a major and a minor groove with

10 base pairs per turn. The A-form is more compact with 11

base pairs per turn. It is observed when DNA is complexed

with RNA, due to the presence of 2'hydroxyl groups on the

ribose preventing formation of the B-form during

complexation. The A-form is also seen under conditions of

low humidity (<75%) or high salt concentration (Umrania et

al. 1995; La Vere et al. 1996). This A-form secondary





37


structure is therefore observed under lyophilized

conditions. While the A and B-forms are both right handed

helixes, the Z-form is a left handed helix and has the most

base pairs per turn of any of the forms and only one

groove. This form has been seen under high salt

concentrations and probably does not form in vivo but could

potentially be seen in low concentrations under the high

salt conditions seen with lyophilization. The secondary

structure of the DNA may effect stability and

functionality.

Lyophilization causes the removal of the hydration

sphere around a molecule. For DNA, it appears that there

are approximately 20 water molecules per nucleotide pair

bound most tightly to DNA that do not form an ice-like

structure upon low temperature cooling. Upon DNA

dehydration at 0% relative humidity only five or six water

molecules remain (Falk et al. 1963; Lith et al. 1986; Tao,

and Lindsay 1989). Lyophilization may increase the

stability of DNA under long-term storage because of

decreased water activity, but may also cause some damage

upon the initial lyophilization process, potentially

through changes in the DNA secondary structure. Agents

that can substitute for nonfreezable water, such as

trehalose, can demonstrate cryoprotective properties for






38


DNA and other molecules during lyophilization of intact

bacteria (Rudolph et al. 1986; Israeli et al. 1993). Other

cryoprotective agents, such as polyols, amino acids, sugars

and lyotropic salts are preferentially excluded from

contact with protein surfaces but are also capable of

stabilizing enzymes during lyophilization by undefined

mechanisms (Carpenter, and Crowe 1989). It is possible

that agents that act as cryoprotectants to proteins may

also act to stabilize nucleic acids.


Materials and Methods


Plasmid Purification

Both pGL3 plasmid and pRL-CMV (Promega, Madison, WI),

respectively encoding for photinus luciferase and renilla

luciferase, were grown in E. coli JM109 cells (Promega,

Madison, WI). The transformed bacteria were cultivated in

Lura Bertina (LB) broth containing 100 pg/ml ampicillin to

select for P-lactamase encoding plasmid. The plasmids were

isolated via an alkaline lysis method and purified using a

silica slurry column (Wizard Plus Megapreps, Promega,

Madison, WI). The plasmid was stored in TE buffer (10 mM

Tris-HCl, 1 mM EDTA, pH 7.4). Plasmid, pRL-CMV, that was to

be lyophilized was aliquoted (50 ul) into 1.5 ml






39


polypropylene tubes (Sarstadt) and frozen at -800C. The

frozen samples were then lyophilized for 24 hours under

vacuum using a Savant Centrivap and a -1050C cold trap.

Lyophilized samples were rehydrated with distilled water

immediately before use.


Liposome Preparation

Lipids were obtained from Avanti Polar Lipids

(Alabaster, AL). Cationic DOTAP : DOPE (dioleoyl glycero

trimethylammonium propane : dioleoyl glycero

phosphoethanolamine) liposomes were prepared using the

hand-shaking method (New 1990). DOTAP: DOPE (10 mg DOTAP,

10 mg DOPE) was added to 10 ml of chloroform and introduced

into a 250 ml round bottom flask. The chloroform was

evaporated using a rotary evaporator at 600C. The lipid

film was hydrated with 10 ml of distilled water and shaken

for 30 minutes at 600C. Lipids were sized by extrusion 6

times through 600 nm polycarbonate filters (Poretics Corp,

Livermore, CA). Sizes were confirmed with a laser light

scattering particle sizer using volume-weighted

distribution (Nicomp 380ZLS, Santa Barbara, CA).





40


Transfection Efficiency Assay

Untreated pGL3 (0.05 pg/ml) and treated pRL-CMV (0.5

pg/ml) were combined with cationic lipid mixture (DOTAP :

DOPE 1:1, 2 pg/ml each) and incubated for 15 minutes to

allow interaction of the anionic plasmid with the cationic

liposomes in TE buffer (Tris HC1, 10 mM; EDTA, 1 mM, pH

7.4). The mixtures were then co-transfected into COS-1

cells (ATCC, Rockville, MD) that had been plated 24 hours

earlier at 3x104 cells/ml in 24 well tissue-culture dishes.

All transfections were effected using serum free DMEM

(Dulbecco's Modified Eagle's Medium) supplemented with

penicillin G (100 units/ml) and streptomycin (100 gg/ml).

The cells were subsequently incubated for 5 hours in a

humidified, 5% CO2 incubator at 370C. The media was

aspirated and the cells washed with PBS (phosphate buffered

saline) before being replaced by DMEM supplemented with 10%

FBS (Fetal Bovine Serum), penicillin G (100 units/ml) and

streptomycin (100 pg/ml). The cells were incubated for an

additional 19 hours, washed with 1 ml PBS, and then lysed

with Passive Lysis buffer (200 pL, Promega, Madison, WI).

Photinus luciferase and renilla luciferase activities

were determined by analyzing the 5 pL of the lysate using

the Dual Luciferase Assay kit (Promega, Madison, WI) and a






41


luminescence spectrophotometer (Monolight 2010, Analytical

Luminescence Laboratory, San Diego, CA). The light output

was measured for 10 seconds and the results integrated to

yield the activity. All pRL-CMV, renilla luciferase

plasmid, measurements were standardized to the ratio of

renilla luciferase (rluc treated) activity to photinus

luciferase (luc control) activity. Essentially, the ratio

of rluc/luc is linear to the concentration of rluc added to

COS-1 cells. Therefore, a linear regression of rluc/luc

ratio at various concentrations of rluc and inactivated

rluc (cut with restriction enzymes), with luc held

constant, allows a determination of the apparent

concentration of the treated pRL-CMV plasmid.


Differential Scanning Calorimetry Analysis

Salmon sperm DNA (Sigma Chemical Company, St. Louis,

MO), was lyophilized in the presence of lyoprotectant (1:1

w/w). Aliquots of 12-15 mg DNA were then placed in crimped

sample pans and the melting profiles determined using a

Seiko 220C Differential Scanning Calorimeter (Tokyo,

Japan). Scans were carried out over a -100C to 1500C

temperature range at a heating rate of 50 per minute.

Nitrogen was used as a purge gas at a flow rate of 100

ml/min. Only a single scan could be accomplished per






42


sample, due to thermal degradation of the DNA/carbohydrate

mixtures. Onset of melt was determined using the

manufacturers software.


DMED Assay

Treated pRL-CMV DNA was incubated with N,N'-

dimethylethylenediamine (100 mM, pH 7.4, 30 minutes at

370C), cleaving abasic sites via p-elimination (McHugh, and

Knowland 1995). The resultant DNA was then visualized

using a 0.8% agarose gel in TBE buffer (50 mM Tris, 50 mM

borate, 1 mM EDTA, pH 7.8). The gel was electrophoresed at

1 V/cm for 16 hours to affect good separation between the

supercoiled, relaxed circular and linear forms of the

plasmid DNA. The gel was then stained with ethidium

bromide (0.5 gg/ml) and photographed on a UV light box using

a DC40 digital camera (Kodak, Rochester, NY).


Analysis of Hyperchromic Effect

Hyperchromic effect was examined using a UV/Vis

spectrophotometer at 260 nm (Lambda 3, Perkin Elmer).

Analysis consisted of comparing the change in absorbance at

260 nm between lyophilized and non-lyophilized samples at

the same 1 mg/ml concentration of plasmid DNA.





43


Oxidative Analysis

Oxidative analysis of plasmid DNA was determined using

an isocratic HPLC method. Plasmid DNA, 100 gg, was first

chemically hydrolyzed in 1 ml of 60% formic acid at 1500C

under vacuum for 45 minutes. Samples were then lyophilized

to remove all formic acid and rehydrated in 2 ml of mobile

phase (50 mM sodium acetate, 1 mM EDTA, 2% methanol, pH

5.5). The samples were then analyzed by HPLC. The HPLC

conditions consisted of a 50 il loop, a 0.8 ml/min flow

rate, and a 270C column temperature. The column used was a

reverse phase C18 column, 25 cm, 5 pm (Microsorb MV, Rainin,

Woburn MA), with a C18 guard column. Detection was

accomplished using a BAS UV-8 detector (West Lafayette, IN)

at 254 nm for non-oxidized bases and a BAS LC-4B

amperometric detector with a range of 10 nA at 750 mV,

using a Ag/AgCl electrode with a glassy carbon working

electrode, for oxidized bases.

Control oxidized DNA was prepared using a Fenton type

iron-catalyzed reaction. Ferric chloride (1 nm) and

hydrogen peroxide (3%) were incubated with pRL-CMV plasmid

(final concentration 1 gg/pl) on ice for 15 minutes. The

plasmid was then purified by ultrafiltration using a 1000

MWCO spin filter (Amicon).






44


Circular Dichroism Analysis

Circular dichroism was analyzed using a Jacso J-500C

spectrapolarimeter (Easton, MD). Scans were performed

using a 1 nm bandwidth, automatic slit-width, 50 mdeg/fs

sensitivity, at 20 nm/min, over a 300-250 nm wavelength

range. All scans were baseline subtracted against DI water

and each data point was reported as the average of at least

two scans. Plasmid DNA conformational change was observed

at 270 nm. Samples were analyzed at 0.5 mg/ml pRL-CMV

plasmid DNA in a 1 cm cuvette, comparing various

weight/weight ratios of pRL-CMV plasmid DNA to

lyoprotectant. Lyophilized samples were rehydrated

immediately before analysis.

Statistical Analysis

Statistical analysis between the various treatments

was conducted using analysis of variance and Fisher's

protected least significant difference (PLSD) post-hoc T-

tests where appropriate (StatView v4.5, Abacus Concepts,

Berkley, CA), with p<0.05 considered statistically

significant.


Results and Discussion

A biofunctional assay was devised to detect damage due

to lyophilization of plasmid DNA. This method is a






45


variation on the conventional cell transfection method,

using two plasmids rather than one. By cotransfecting both

control and treated plasmid the variation in transfection

efficiencies can be taken into account. The treatment

plasmid was the pRL-CMV construct (Promega, Madison, WI),

containing the renilla luciferase gene driven by a

cytomegalovirus promoter. This version of luciferase was

isolated from the sea pansy, a species of bioluminescent

coral. The control plasmid was the pGL3 construct

(Promega). The construct encodes the photinus luciferase

gene driven by a simian virus promoter. The two versions

of luciferase each require different cofactors and can be

assayed in the same test tube, since the conditions

appropriate for the renilla luciferase reaction quench the

photinus luciferase reaction. The two plasmid constructs

were transfected into COS1 cells, an African green monkey

kidney cell line, that expresses the large T antigen

transcription factor needed to drive the SV40 promoter.

This transcription factor was capable of upregulating the

CMV promoter, as well (Soneoka et al. 1995). The

transfections were liposome mediated with the optimal ratio

of lipid to pRL-CMV to pGL3 for detection as suggested by

the manufacturer. This ratio was empirically based upon

the linearity of concentration to activity standard curves






46



(Figure 3) and the need to prevent cross talk between the


different promoters present in the two vectors. The cells


were transfected in serum free media and were incubated for


,0.7- ---- 0.7
0.6- 0.6
0.5 0.5
E0.4 E 0.4
< 0.3 40.3
z *
S0.2 0 02
0.1- 0.1
0 0
Cntrd 4 2 1 0.1 Cntd 4 2 1 0.1
Glucose:DNA (wlw) Glucose-l-Phosphate:DNA (wtw)



0.7 0.7-
0.6 0.60

E 0.5 2 0.4
E 0.4- E 0.4 -
0.3 0.3
S0.2 00.2 *
0.1- M 0 0.1

Cntri 4 2 1 0.1 Cntr 4 2 1 0.1
Glucosamine:DNA (wlw) Sucrose:DNA (wfw)



0.7- 0.7
0.6 0.6



0.5 0.1
0o.l I0 i I A

0 I .0.0
Cntrl 4 2 1 0.1 Cntr 4 2 1 0.1
Lactose:DNA (wlw) Urea:DNA (w/w)

Figure 8: Effect of lyophilization (FD) on plasmid DNA
activity, with various amounts lyoprotectant to DNA (w/w).
Average + SEM. n=5 for all treatments, = FD DNA
significantly lower than Control DNA. p<0.05 via Schefe's
multiple comparison T-test.


5 hours. The media was then replaced with serum containing


media and incubated a further 48 hours before being lysed.






47


Both forms of luciferase were then assayed using a

luminescence spectrophotometer (Monolight 2010).

The biofunctionality assay demonstrated a loss of more

than 75% of plasmid DNA activity after lyophilization

(Figure 8). Furthermore, this loss of activity could be

prevented by the use of lyoprotectant carbohydrates

(approximately 4:1 carbohydrate to DNA w/w), including

glucose, glucose-1-phosphate, glucosamine, lactose and

sucrose). These carbohydrates were chosen to study the

effect of charge and size on the effectiveness of the

lyoprotection.

The presence of lyoprotectant did not significantly

alter the apparent plasmid DNA concentration either before

(data not shown) or after freeze-drying. The protective

effect of the glucose-1-phosphate was unexpected if the

lyoprotective effect was due to the replacement of the

sphere of hydration by the carbohydrate, as it might be

assumed that this anionic carbohydrate would be

electrostatically repulsed from the major groove by the

anionic phosphate backbone. Furthermore, glucosamine,

which would be expected to interact strongly with plasmid

DNA, did not exert a statistically significant protective

effect at a 4:1 w/w ratio. If the lyoprotectant

carbohydrates are protective of plasmid DNA due to






48


replacement of the water sphere of hydration, then the use

of a water destabilizing compound such as urea should

decrease DNA activity. However, as seen in Figure 8, urea

did support a protective effect through an undetermined

mechanism.

Disaccharides demonstrated a greater protective effect

on a molar basis than did monosaccharides, suggesting that

the size of the protective compound may exert and

influence. Potentially, the effect could be due to a non-

interactive covering of the plasmid DNA that inhibits any

conformationally induced denaturation. This hypothesis is

supported by the protective effect seen upon "saturating"

concentrations of all lyoprotectants tested (4000:1 w/w).

The extent of interaction between DNA and the

lyoprotectant carbohydrates was assessed using thermal

analysis to determine onset of melting temperatures.

First, behavior in the solid phase was examined using

differential scanning calorimetry, a sample DSC scan can be

seen in Figure 9. Equal weights of carbohydrate and

salmon-sperm DNA were mixed and lyophilized. The

lyophilized cake was then assayed to determine the onset of

melting. Second, the effect of lyophilization upon melting

temperature after rehydration was analyzed using UV

spectroscopy. In the dried state, there was a general






49


trend towards increased onset of melting temperature in the

presence of lyoprotectants Figure 10, with a change in the

onset of melting suggesting an interaction between the

carbohydrates and the DNA. However, glucose-l-phosphate

did not show a significant increase in Tm, but did protect

against loss of bioactivity, which may suggest that a

physical interaction is not necessary for a protective

effect.


5





51.4 C -
12.68 iln
-3. O0 -




-1



-10
-7.S\ -10





1G-I .l5 -15
-11.7 5.8 23.3 40.8 58.3 75.8 93.3 110.8 128.3 145.8
University of Flortil-MSE TEMP C (Heating)



Figure 9: Representative thermal analysis scan of salmon
sperm DNA.


The effect of rehydration of lyophilized plasmid DNA

on the DNA melting profile, as measured via absorbance at

260 nm, can be seen in Figure 11. Lyophilized-rehydrated






50


plasmid DNA showed an earlier onset of melting than plasmid

DNA that was kept in solution. This appears to agree with

the lower onset of melting seen in the solid state without

lyoprotection. It can be seen that the previously

lyophilized DNA solutions showed an increased absorption

over the 25-800C interval, indicative of the presence of a

small amount of single-stranded DNA in the DNA solutions

(Lindhal and Nyberg 1972).

80


60


E 40


20


0





Figure 10: Effect of lyoprotectants on the melting
temperature of salmon-sperm DNA. Mean + SEM, n=3, =
significantly different than unprotected control DNA (p <
0.05) via Fisher's PLSD.

To identify a potential mechanistic explanation for

the results obtained by the biofunctionality assay, several

structural assays were also utilized. Standard agarose gel

electrophoresis (0.8% agarose) is capable of separating the

conformations of plasmid such as supercoiled, relaxed






51


circular and linear, by utilizing size fractionation in the

agarose gel via an electric field. The DNA is negatively

charged because of its phosphate backbone, and will migrate

toward the cathode when an electric field is applied. No

apparent change in the ratio of supercoiled to relaxed

plasmid DNA was observed using this method, Figure 12 and

Figure 13, suggesting that the lyophilization process did

not cause any gross conformational changes to the plasmid

DNA.

0.6

0.5

0.4
v i-FD
N 0.3
.3 J-g-Sin

0.2

0.1

0.0
o.o -M fs 9 '-------- -
25.0 35.0 45.0 55.0 65.0 75.0 85.0
Temperature (C)
Figure 11: Effect of lyophilization and subsequent
rehydration on the melting of plasmid DNA. Mean + SEM, n=3

The standard agarose gel method can not differentiate

between undamaged supercoiled and damaged supercoiled

plasmid DNA. This damaged supercoiled plasmid DNA could

potentially be seen after glycosidic bond cleavage at






52




















Figure 12: Lyophilized plasmid DNA samples, rehydrated in
DI water and run an agarose gel. Samples: marker, HindII
digested lambda phage marker; FD, lyophilized DNA without
DMED treatment; FD DMED, lyophilized DNA with DMED
treatment; pH 3, DNA incubated in pH 3.0, 2.5 mM citrate
buffer for 15 minutes at 500C with no DMED treatment; pH 3
DMED, DNA incubated in pH 3.0, 2.5 mM citrate buffer for 15
minutes at 500C with DMED treatment before running on gel.

abasic sites. Abasic sites are one of the most common DNA

lesions and can be produced in at least two ways (McHugh,

and Knowland 1995): via spontaneous hydrolysis of the

glycosyl bond between deoxyribose and the purines, or at a

slower rate between deoxyribose and pyrimidines. These

modifications can be accelerated by base modifications such

as the alkylation of purines, saturation of C5-C6 bond of

the pyrimidines, and fractionation of the heterocyclic ring

(Talpaert-Borle, and Liuzzi 1983). In addition, they can

be detected by the use of the DMED assay, which cleaves the






53


phosphate backbone by a beta elimination reaction in the

presence of an abasic site (Figure 2 on page 7).

100%


80%


60%
o
S 40%


20%


0%
0 1 2 3
Lyophilization Cycles
Figure 13: Effect of lyophilization on plasmid form. Mean +
SEM, n=3. p > 0.05 via ANOVA

After incubation in the presence of DMED damaged DNA

would be expected to show a lower percentage of supercoiled

DNA. The percentage of supercoiled DNA did not change

after lyophilization and subsequent DMED treatment (Figure

12). This suggests that the lyophilization process did not

cause an increase in plasmid DNA abasic sites.

Analysis by UV/Vis spectroscopy showed a hyperchromic

effect for lyophilized plasmid DNA, suggesting potential

denaturation or conformational change. Any process which

increases the interaction of purine and pyrimidine rings,

such as a contraction of the plasmid DNA macromolecule or






54


restriction of internucleotide rotation by the formation of

hydrogen bond stabilized helical structures, would result

in a hyperchromic effect at 260 nm, as would separation of

the two anti-parallel strands (denaturation) (Michelson,

1963). Lyophilization of plasmid DNA resulted in a

significant increase in absorbance at 260 nm. This effect

can be alleviated by the use of lyoprotectant carbohydrates

at a 50 mM concentration (Figure 14). These results

suggested that the lyophilization process causes a

conformational change or denaturation in plasmid DNA that

can be prevented by the use of carbohydrates as

lyoprotectants, since these data agree with the UV melting

profile which also suggests a denaturation.

20


O 15
0
x
E
o 10






0




Figure 14: Effect of lyophilization on plasmid DNA
absorbance at 260 nm. Average + SEM. n=4, FD DNA
significantly higher than lyoprotected DNA p<0.05 via ANOVA






55


To further identify possible conformational changes

circular dichroism analysis was utilized. Lyophilized

plasmid DNA was compared to "normal" B-form DNA (pH 7.4 TE

buffer) and to high-salt, A-form DNA (5 M NaC1) (Bailleal

et al. 1984; Nishimura et al. 1985). As can be seen in

Figure 15, the high salt conditions seen during

lyophilization cause a conformational change that may be

indicative of a change from the B-form to the A-form.



6

?5 FD

4 A




1

0
250 260 270 280 290 300
Wavelength (nm)


Figure 15: Wavelength circular dichroism scan of
lyophilized plasmid DNA (FD) compared to A-form and B-form
plasmid DNA. Each scan is the average of three separate
baseline subtracted scans.

The use of lyoprotectants can exert further

conformational changes during the lyophilization process

Figure 16. The smallest uncharged protectant, glucose,

appeared to undergo the least conformational change during

the lyophilization process. Increasing the size of the





56

protective molecule, by utilizing lactose as a protectant

causes a slight decrease in conformational shift, but not

nearly as great a shift as the smaller glucose monomer.
When charged moieties were added to the glucose protectant,

a further shift in conformation was noted. Both anionic

and cationic compounds caused a shift in conformation

a 3.0 3.0
2.5- A g 25 B
2. I I 20I


tI 1 111 1. 1 I
to is






0s5 05 Df





Figure 16: Effect of lyoprotection on plasmid DNA
conformational change. Panels each signify a separate
protectant; A: lactose, B: glucosamine, C: glucose-1-
phosphate, and D: glucose. For each panel Protectant:DNA
1, 5, 10, 20 and 40 or 1, 2, and 8 w/w ratios. Cntrl S
"protectant" is protectant:DNA solution (40:1 w/w) non
lyophilized. Cntrl FD and Cntrl S "without protectant" are
no protectant, lyophilzed or non lyophilized respectively.
n=10, mean + SEM.

towards the A-form, with the anionic glucose-1-phosphate
having a two-fold increase in CD shift over the cationic
glucosamine, causing a conformational change even in the
glucosamine, causing a conformational change even in the





57


solution control state. Furthermore, there is a lack of a

clear dose-response relationship between the amount of the

carbohydrate and the conformational shift for all compounds

except glucose-l-phosphate. It seems that the anionic

glucose-l-phosphate requires a 5:1 (w/w) protectant:DNA

ratio before the protectant exerts a greater conformational

change upon the DNA. This increase in CD measurement is

typical of a change in both the helix winding angle and the

base pair twist of the plasmid DNA. An increase in the CD

measurement signals a shift towards the A-form of the

plasmid DNA, with an a decreased winding angle, angle

between two adjacent bases, leading to more base pairs per

turn of the double helix (Johnson et al. 1981).

The kinetics of the conformational change were

examined using CD. Essentially, DNA was lyophilized and

consequently rehydrated. CD spectra were recorded for

multiple time points after rehydration (Figure 17). A

first order fit of the data suggests that most of the

plasmid returns to the B-form in approximately five hours.

At 24 hours after rehydration the measured CD was not

statistically different from control non-lyophilized DNA

(data not shown), this suggests that the conformation of

the plasmid DNA may return to the B-form within 24 hours.






58


It was hypothesized that this initial conformational

change could be responsible for the apparent loss of

plasmid DNA activity after lyophilization. Since it was

shown that the plasmid returns to the B-form in a

relatively short period of time, plasmid DNA was

lyophilized and rehydrated immediately before transfection

or rehydrated 24 hours before transfection to determine the

A-form effect on plasmid DNA activity (Figure 18). There

was no significant difference between 0 and 24 hours

rehydration, suggesting that this potential conformation of

the plasmid DNA does not have a direct impact on biological

activity.


0.8
0.7
0 0.6
0.5 -
0.4
0.3 -
S0.2

0.1
0.0 ,
0.00 0.08 0.16 0.25 0.50 1.00 24.00
Hours Rehydrated



Figure 17: Kinetics of plasmid conformational change after
rehydration. Mean + SEM, n=3.






59


Another mechanism of damage may be oxidative damage of

the plasmid DNA during the lyophilization process. To

examine the possibility of this mechanism, plasmid DNA was

lyophilized up to six times and then chemically hydrolyzed

to the individual bases. The bases were then analyzed by


0.7
0.6
0.5 -
c 0.4
E
< 0.3 -
0.2
0.1 -
0.0
0 24 Cntrl
Rehydration Tine (Hours)



Figure 18: Effect of rehydration time on plasmid DNA
activity. Mean + SEM. n=6. No significant difference
between 0 and 24 hours by T-test.

HPLC using parallel UV and ECD detection to quantitate both

normal and oxidized bases, specifically guanine and 8-OH-

guanine. The 8-OH-guanine is a commonly used marker for

oxidative damage of DNA. The lyophilized samples showed no

significant increase in 8-OH-guanine per mole of guanine

and all lyophilized samples were significantly less damaged

than the positive control (Figure 20).

The type of tube used for lyophilization of plasmid

DNA could potentially change the biological activity of





60


plasmid DNA (Figure 21). If the loss of biological

activity was due to the interaction of plasmid DNA and the

container used for lyophilization, then the use of





A B





\l







Figure 19: HPLC analysis of hydrolyzed pRL-CMV plasmid DNA.
Panel A is a UV analysis. Peaks by retention time: 2.76,
solvent front, 3.79 cytosine, 7.85 guanine, 11.15 thymine,
13.00 unidentified, 17.89 adenine. Panel B is an ECD
analysis. Peaks by retention time 2,67 3.21 solvent
front, 3.53 5-OH cytosine, 3.82 cytosine, 4.78 plasticizer,
7.35 guanine, 7.92 8-OH guanine, 16.99 8-OH adenine and
adenine.

siliconized tubes would be expected to decrease DNA loss.

However, there is no significant difference in DNA activity

between siliconized and non-siliconized polypropylene

tubes, suggesting that plasmid DNA does not interact with

the polypropylene tube during the lyophilization process.

This is supported by gel electrophoresis, which showed no

significant loss of plasmid DNA after cycles of






61


140

0 120

E 100
C
o 80



040
o*
E 20 *


0 2 4 6 Cntrl
Freeze-drying Cycles

Figure 20: Effect of lyophilization of plasmid DNA on
oxidative damage. Positive control was Fe+3 catalyzed
oxidation. Mean + SEM, n=5, except control n=3. = p <
0.05 via Fisher's PLSD versus control.

lyophilization. There did appear to be a significant

interaction between polyethylene and plasmid DNA during the

lyophilization process, suggesting that polyethylene tubes

are not an appropriate choice for storage of plasmid DNA.


Conclusion

In conclusion, lyophilization of plasmid DNA causes a

loss of plasmid DNA functionality that can be prevented by

the use of carbohydrates as lyoprotectants. The mechanism

behind this loss of activity does not appear to be due to a

gross structural change, as would be evidenced via agarose

gel electrophoresis. Nor does the damage appear to be due

to an increase in plasmid DNA abasic sites, as measured by





62


the DMED assay, increase in oxidative damage, as measured

via HPLC, or interaction with the enclosure used for


E 0.7-

-0.6

0 0.5
< 0.4
S*
P.0.3

0.2
"X
10.1


Cntri PP SP PE
Figure 21: Effect of tube type on plasmid DNA bioactivity
after lyophilization. Cntrl = non-lyophilized DNA, PP =
polypropylene, SP = siliconized polypropylene, PE =
polyethylene. Mean + SEM, n=5, = p < 0.05 via Fisher's
PLSD versus control, X = p < 0.05 versus PP and SP.

lyophilization. However, this change in plasmid DNA

activity does correlate to a change in plasmid hyperchromic

effect, as evidenced by a change in absorbance at 260 nm,

and a change in the melting temperature, as evidenced by

DSC and UV measurements. These data suggest that plasmid

DNA damage after lyophilization is mediated by some sort of

conformational change but the conformational of the plasmid

DNA does not directly affect plasmid activity, as suggested

by circular dichroism spectra. This conformational change

is not prevented by all carbohydrates tested, but all





63


protectants were effective against the hyperchromic effect.

This suggests that lyoprotectants may decrease a

conformationally-induced denaturation of the plasmid DNA

caused by the lyophilization process.















CHAPTER 4
CHARACTERIZATION OF ENDOTOXIN AND CATIONIC LIPOSOME
INTERACTION


Introduction

Gene therapy is one of the fastest growing areas in

therapeutics. While rapid progress has occurred in this

field, the pharmaceutical community has not adequately

addressed safety concerns. In particular, there exists a

potential toxicity from plasmid DNA contaminates that could

be exacerbated by conventional non-viral gene delivery

methods. Since plasmid DNA is typically produced by gram-

negative bacteria E. coli, endogenous bacterial products

can potentially contaminate the final plasmid DNA product

(Weber et al. 1995). Typical problematic agents in plasmid

DNA preparations include endotoxin (also known as

lipopolysaccharide or LPS), a cell wall component that is

pyrogenic in man, and DNase, a ubiquitous bacterial enzyme

that can degrade the final plasmid product and act as an

immunogen.

Endotoxin ranks among the most serious limitations in

the development of gene products for gene therapy because



64





65


of its extreme toxicity. E. coli endotoxin consists of a

polysaccharide component and a covalently bound lipid

component, lipid A. Lipid A is the biologically active

component responsible for pathophysiological effects

including fever, hypotension, intravascular coagulation and

potentially death (Fletcher, and Ramwell 1980; Aida, and

Pabst 1990; Rietschel et al. 1993; Xing et al. 1994).

Endotoxin is an extremely potent toxin with an LD5o of 3

mg/kg and 1 mg/kg in rats and dogs, respectively (Fletcher,

and Ramwell 1980; Shibayama et al. 1991). It has been

shown that as little as 150 pg of endotoxin can be lethal to

a horse (Bottoms 1982). Therefore, the Food and Drug

Administration (FDA) has established a guidance on human

maximal endotoxin dose permissible for parenteral products

(F.D.A 1985). This limit is based on endotoxin activity

(EU), and can be measured via the LAL (Limulus amebocyte

lysate) assay (Levin, and Bang 1964; Levin, and Bang 1968;

Iwanaga 1993). Non-intrathecal parenteral drug products

are limited to 5 EU kg-1 hr-1. In the case of an intrathecal

parenteral product, the injection limit drops to 0.2 EU kg-1

hr-1. These allowable levels are of concern in gene

therapy, where the extent of endotoxin contamination in

plasmid DNA preparations after treatment to remove

endotoxin has been shown to range from 3-15 EU/mg plasmid





66


DNA, with 1 EU = 1 ng pure endotoxin (Cotton et al. 1994).

Determinations in our laboratory have shown that before

specific removal, endotoxin contamination of plasmid DNA

can typically approach 5,000 EU/mg, while the literature

reports endotoxin concentrations exceeding 15,000 EU/mg DNA

(Montbriand, and Malone 1996).

It was previously shown that increasing levels of

endotoxin contamination decrease transfection efficiency

when using DOTAP : DOPE (dioleoyl glycero trimethylammonium

propane : dioleoyl glycero phosphoethanolamine) cationic

liposomes (Weber et al. 1995) or adenovirus particles

(Cotton et al. 1994) to deliver plasmid DNA. However, the

mechanism underlying this decreased transfection efficiency

has not been clearly defined. It can be predicted that

endotoxin interacts with cationic liposomes

electrostatically due to the positive charges on the

cationic liposomes and the negatively charged

phosphorylated glucosamine residues of the lipid A moiety

(Rietschel et al. 1993). There also may be other

lipophilic methods of interaction with cationic lipids due

to the saturated fatty acid chains on lipid A (Rietschel et

al. 1993). Upon this interaction of endotoxin, DNA and

cationic liposomes, the liposome complex would be expected

to be taken into the cell via endocytosis with subsequent






67


destabilization of the endosome and release of the DNA and

endotoxin into the cytoplasm (Xu, and Szoka 1996).

It will be of importance for formulation scientists to

understand the potential modes of interaction between

endotoxin and cationic liposomes. With this knowledge it

should be possible to develop newer methods of endotoxin

removal from these products. In this report we have

focused on endotoxin, similar concerns could be raised with

other anionic compounds.


Materials and Methods


Plasmid Purification

A pGL3 plasmid (Promega, Madison, WI) encoding for

luciferase driven by the SV-40 promoter was propagated in

E. coli JM109 cells (Promega, Madison, WI). The

transformed bacteria were cultivated in Lura Bertina (LB)

broth containing 100 gg/ml ampicillin to select for 0-

lactamase encoding plasmid. The pGL3 plasmid was isolated

via an alkaline lysis method and purified using an anionic

exchange column (Qiagen, Chatsworth, CA). The plasmid was

stored in TE buffer (Tris HC1, 10 mM; EDTA, 1 mM, pH 7.4).

Endotoxin content of the plasmid DNA before endotoxin

removal was approximately 4,000 EU/mg, as determined by the

LAL assay method (QCL-1000, BioWhittaker, Walkersville,





68


MD). Endotoxin was removed by the Triton X-114 method of

Manthorpe, et al (Manthorpe et al. 1993).


Liposome Preparation

Lipids were obtained from Avanti Polar Lipids

(Alabaster, AL). Cationic DOTAP : DOPE, NBD labeled DOTAP

: DOPE, and zwitterionic lecithin liposomes were prepared

using the hand-shaking method (New 1990). DOTAP : DOPE (10

mg DOTAP, 10 mg DOPE), NBD labeled DOTAP : DOPE (10 mg

DOTAP, 10 mg DOPE, 0.1 mg l-palmitoyl-2-[12-[(7-nitro-2-

1,3-benzodiamino]dodecanoyl]-sn-glycero-3-phosphate), or

lecithin (10 mg) was added to 10 ml of chloroform and

introduced into a 250 ml round bottom flask. Chloroform

was evaporated using a rotary evaporator at 600C. The lipid

film was hydrated with 10 ml of distilled water and shaken

for 30 minutes at 600C. Lipids were sized by extrusion 6

times through 200, 600, or 800 nm polycarbonate filters

(Poretics Corp, Livermore, CA). Sizes were confirmed with

a laser light scattering particle sizer using volume-

weighted distribution (Nicomp 380ZLS, Santa Barbara, CA).


Transfection Efficiency Assay

Phenol purified, unlabeled endotoxin (0-50,000 EU/ml),

E. coli serotype 055:B5 (500 EU/.g, Sigma Chemical Company,






69


St. Louis, MO) was added to a pGL3 luciferase plasmid :

cationic lipid mixture (DOTAP : DOPE 1:1 w/w) with a ratio

of 1 gg plasmid to 2 gg DOTAP. The mixtures were incubated

for 15 minutes to allow interaction of the anionic plasmid

to the cationic liposomes, followed by transfection into

COS-1 cells (ATCC, Rockville, MD) that had been plated 24

hours earlier at 3x104 cells/ml in 24 well tissue-culture

dishes. All transfections were carried out using serum

free DMEM (Dulbecco's Modified Eagle's Medium) supplemented

with penicillin G (100 units/ml) and streptomycin (100

gg/ml). They were subsequently incubated for 5 hours in a

humidified, 5% CO2 incubator at 370C. The media was

aspirated and the cells washed with PBS (phosphate buffered

saline) before being replaced by 10% FBS (fetal bovine

serum) containing DMEM and antibiotics. The cells were

incubated for an additional 19 hours, washed with 1 ml PBS,

and then lysed with lysis buffer (200 gL, 0.1 M potassium

phosphate, 1% Triton X-100, 1 mM DTT, 2 mM EDTA, pH 7.8).

Luciferase activity was determined using a

luminescence spectrophotometer (Monolight 2010, Analytical

Luminescence Laboratory, San Diego, CA). Lysate (20 iL) and

the assay buffer (100 iL, 30 mM tricine, 3 mM ATP, 15 mM

MgSO4, 10 mM DTT, pH 7.8) were added to the sample cuvette.






70


D-luciferin (100 pL, 1 mM, pH 6.5) was injected to initiate

the reaction. The light output was measured for 10 seconds

and the results integrated to yield the luciferase

activity. All luciferase measurements were standardized

against protein concentration as measured by the

bicinchoninic acid protein assay (BCA Protein Assay,

Pierce, Rockford, IL).


Anisotropy Assay

A constant amount (1000 EU) of flourescein

isothiocyanate (FITC) labeled endotoxin, E. coli serotype

055:B5 (100 EU/pg, Sigma Chemical Company, St. Louis, MO),

and varying amounts of unlabeled cationic DOTAP : DOPE

liposomes (600 nm, 0-16 ig), zwitterionic lecithin liposomes

(200 and 800 nm, 0-16 pg,), PAMAM (polyamidoamine)

dendrimer (generation 4, 0-16 ig, in TE buffer, pH 7.4,

Aldrich Chemical, Milwaukee, WI), or plasmid DNA (pGL3 in

DI water), with a final volume of 1 ml, were allowed to

equilibrate together for 15 minutes at 250C. Fluorescent

anisotropic measurements were conducted at 250C, 487 nm and

525 nm excitation and emission wavelengths, respectively

using a luminescence spectrophotometer (LS50B, Perkin

Elmer, Oak Brook, IL).






71


Endotoxin Dephosphorylation

Unlabeled endotoxin (30 EU) was incubated for various

time from 0 to 120 minutes with 1 mU alkaline phosphatase

(Promega, Madison, WI) at 370C following the methods of

Poelstra, et al (Poelstra et al. 1997a; Poelstra et al.

1997b) with minor variation in the assay buffer: 0.5 M

Tris, 10 mM MgCl2, 1 mM ZnCl2, pH 7.8.


Endotoxin Assay

Endotoxin activity was quantitatively measured using a

commercially available chromogenic LAL test method (QCL-

1000, BioWhittaker, Walkersville, MD) according to the

manufacture's instructions. In brief, endotoxin (0.6 EU)

was incubated for 15 minutes with an increasing

concentration of DOTAP: DOPE cationic liposomes (0-100 ng)

and brought to a final volume of 50 gL with endotoxin free

water. The LAL reagent (50 pL) was added to the reaction

vessel and the mixture was further incubated for 10 minutes

at 370C. Chromogenic substrate solution (100 gL) was added

and the sample incubated for 6 minutes before the addition

of stop reagent (100 RL, 25% glacial acetic acid in water).

Absorbance of the p-nitroanaline product was read at 405 nm

using a UV/Vis spectrophotometer (Lambda 3, Perkin Elmer,

Oak Brook, IL)






72


Cell Viability Assay

The MTT (dimethylthiazol diphenyltetrazolium bromide)

assay (Freshney 1994) was used to assess the effect of the

endotoxin and liposome delivery systems on cell viability.

COS-1 cells were plated in a 24 well tissue-culture plate

at a density of 3x104 cells/well in 1 ml DMEM media

containing 10% FBS and incubated for 12 hours in a 370C,

humidified, 5% CO2, incubator. The serum containing media

was aspirated, washed with PBS, and replaced with serum

free media. Varying amounts of lipid (0-170 gg), endotoxin

(0-50,000 EU), and lipid: endotoxin combinations (5 pg : 0-

50,000 EU) were added to each well and incubated for 5

hours. The media was then changed back to 10% FBS DMEM and

the incubation continued until 24 hours following the

addition of test formulations. The cells were then fed

with 1 ml of fresh media and MTT (250 pL, 5 mg/ml) and were

incubated for 5 hours and then the media was removed.

Dimethylsulfoxide (DMSO, 1 ml) and glycine buffer (250 pL,

0.1 M glycine, 0.1 M NaC1, pH 10.5) were added and the

absorbance was immediately read at 570 nm. Untreated cells

and buffer alone were used as positive and negative

controls, respectively.






73


Statistical Analysis

Statistical analysis between the various treatments

was conducted using analysis of variance and Fisher's

protected least significant difference (PLSD) post-hoc T-

tests where appropriate (StatView v4.5, Abacus Concepts,

Berkley, CA), with p<0.05 considered statistically

significant.


Results

The effects of endotoxin contamination upon

transfection efficiency were determined by adding increased

levels of endotoxin to pGL3 luciferase plasmid : cationic

lipid mixture (DOTAP : DOPE 1:1 w/w) with a ratio of 1 pg

plasmid to 2 pg DOTAP before transfection into COS-1 cells

(Figure 22). A 1:2 ratio of plasmid to cationic lipid

results in a neutral to slightly negative net charge for

the overall complex. In the presence of DOTAP : DOPE

cationic liposomes (2 pg/ml), low levels of endotoxin (50

EU/ml) appeared to increase the variability of luciferase

reporter activity, a marker of the plasmid DNA

functionality. Higher concentrations of endotoxin resulted

in a corresponding decrease in the luciferase reporter

activity. Activity decreased more than 90% at 5000 EU/ml






74


endotoxin, however the cells appeared viable under direct

observation.

50

40

3c 30
-D Liposome
t-- No Liposome
o)20



0

0 50 500 5,000 50,000
Endotoxin (EU/mL)
Figure 22: Enzyme activity corrected for total cellular
protein after transfection of luciferase plasmid (lpg) in
the presence of endotoxin, with and without DOTAP : DOPE
cationic liposomes. with lipid (2 ug/ml), U without
lipid. RLU+SEM, n=4, p<0.05 via one way ANOVA for effect of
endotoxin in the presence of cationic lipid.

The extent of interaction between endotoxin and

cationic liposomes (DOTAP : DOPE) was determined using

fluorescence anisotropy. The potential contribution of

electrostatic versus lipophilic interactions was

investigated by varying the ionic strength of the

incubation solution (NaC1, 0-2 M). There was a correlative

increase in the anisotropic measurement when endotoxin was

incubated with additional cationic lipid, suggesting

formation of a complex (Figure 23). Maximum binding

occurred at a 1:2 (w/w) ratio of total lipid to endotoxin.






75


The interaction decreased with increasing ionic strength,

but the size of the complex increased significantly, as

monitored by change in anisotropy, even under high ionic

strengths (2 M NaC1), suggesting the presence of

electrostatic in addition to other interactions. Free FITC

was used as a control to ensure the interaction detected

was due to endotoxin and not a consequence of the FITC

label (Figure 23 inset). Furthermore, the spectral

properties of the free FITC and the conjugated FITC appear

similar as determined by excitation and emission maxima.

The potential for competition between plasmid DNA and

endotoxin for DOTAP : DOPE cationic liposome was compared

using a similar experimental paradigm. DOTAP : DOPE (5 gg)

and FITC conjugated endotoxin (1000 EU) were held constant

at the maximal binding ratio seen in Figure 23. As plasmid

DNA levels were increased, there was a decrease in

anisotropy readings, suggesting increased competition

between endotoxin and plasmid DNA for interaction with

DOTAP : DOPE. This results in the displacement of

endotoxin (Figure 24).

To further characterize the mechanism of endotoxin and

cationic liposome interaction, additional fluorescence

anisotropy studies were carried out. Two sizes of PAMAM

dendrimers, a cationic cascade polymer, were studied. The






76


0.12

0.12
0.10- 0.10 FRe Frrc
CL>. o.o0
I 0.06
0.04
0.08- 0.02
o.oo -- DI Water
C. 0 4 8 12 16 -w 0.5 M NaCI

S0.06 -- 2 M NaCI





on anisotropy (r) DI Water, 05 NaC, A 1.0 M NaCI
X 2.0 M NaC. Inset: Free FITC (22 ng 1.0 M NaC. ChangeaCI
0.024



0.002

0in rS2 4 6 8 10 12 14 16
DOTAP:DOPE (PAg)

Figure 23: Effect of increasing DOTAP : DOPE liposomes
concentration with FITC-endotoxin held constant (1000 EU)
on anisotropy (r). DI Water, N 0.5 M NaCl, A 1.0 M NaCl,
X 2.0 M NaCl. Inset: Free FITC (22 ng) 1.0 M NaCl. Change
in r+SEM, n=3, p<0.05 using two way ANOVA for increase in r
with increasing lipid under all conditions.


dendrimers would be expected to interact with FITC labeled

endotoxin though electrostatic forces since the dendrimers

only exposed functional groups are amines. Also, two sizes

of zwitterionic lecithin liposomes (0.2 and 0.8 pm) were

examined to determine the importance of the other

interactions between the lipids. The greatest change in

anisotropy occurred with the cationic PAMAM dendrimers

(Figure 25). Smaller, though statistically significant,

changes in anisotropy were seen at the same concentration

of lecithin, suggesting that the prevalent mechanism of






77






0.12


0.10


0.08


0.06


0.04


0.02-


0.00 I I '- I-' I1 I- I *- I-
0 2 4 6 8 10 12 14 16
pDNA (9g)

Figure 24: Effect of increasing luciferase plasmid
concentration on anisotropy (r) with constant FITC-
endotoxin (1000 EU) and DOTAP : DOPE liposomes (5 gg).
Change in r+SEM, n=3, p<0.05 using two way ANOVA for
decrease in r with increasing plasmid.

interaction is electrostatic rather than lipophilic. In

the case of both sizes of dendrimers and liposomes, the

larger sized particles led to a greater change in

anisotropy as expected, due to the additive nature of

anisotropy. Further evidence for interaction between

endotoxin and cationic liposomes was demonstrated using a

gel retardation assay (Figure 26), which confirmed the





78


FITC-endotoxin cationic lipid complex by retarding the

progress of the endotoxin band in an agarose gel.

0.16

0.14

0.12

S0.10
>-+ Dendrimer G4
o. -- Dendrimer G2
008 -- Uposome 0.8pm
S-*- ULposome 0.2tim
< 0.06

0.04

0.00

0 5 10 15
pg added
Figure 25: Effect of increasing lipid or dendrimer
concentration with FITC-endotoxin held constant (1000 EU)
on anisotropy (r). U Dendrimer (Generation 2), Dendrimer
(Generation 4), X lecithin liposomes (0.2 pm), A lecithin
liposomes (0.8 pm), Change in r+SEM, n=3, p<0.05 using two
way ANOVA for increase in r with increasing concentration
under all conditions.


Well


LPS







0 3 6 8 10 12 14 16
DOTAP:DOPE (lg)

Figure 26: Effect of increasing DOTAP : DOPE liposome
concentration with FITC-Endotoxin held constant (10 Vg).






79


The effect of dephosphorylation of endotoxin on

cationic liposome interaction was also examined (Figure

27). Unlabeled endotoxin (30 EU) was incubated for varying

time with calf intestinal alkaline phosphatase. The

dephosphorylated endotoxin was allowed to interact with

fluorescent NBD labeled DOTAP : DOPE cationic lipid (10 gg,

600 nm). Increased incubation time with the alkaline

phosphatase led to a trend towards decreased anisotropy

(p=0.2 via ANOVA). The anisotropy signal seen after 120

minutes of endotoxin incubation was statistically

equivalent to labeled lipid alone. The small change in

anisotropy was due to the label attached to the liposome

and to the high ionic strength of the reaction buffer,

resulting in a small size change upon interaction.

The chromogenic LAL assay was used to determine the

effect of interaction of endotoxin and cationic lipid upon

the toxicity of the lipid A moiety. All determinations

were compared to a standard curve of known endotoxin

concentrations. No endotoxin activity was evident with the

cationic lipid alone. The LAL assay showed no significant

change in the endotoxin activity when free endotoxin was

compared with cationic lipid-endotoxin complex (Figure 28).






80






0.025


0.020


0.015


S0.010 -


0.005


0.000 I I I II
0 20 40 60 80 100 120
Time (min)


Figure 27: Effect of increasing incubation time of alkaline
phosphatase and unlabeled endotoxin (33 EU) on anisotropy
(r) with constant and NBD labeled DOTAP : DOPE liposome (10
pg). Change in r + SEM, n=4, p>0.05 via one way ANOVA for
decrease in anisotropy over time.

Literature reports suggest that cationic lipids are

toxic in vitro and in vivo (Scheule et al. 1997); however

the toxicity of the cationic lipid endotoxin complex is

not known. The MTT assay was used to determine the effect

of the complex on COS-1 cell viability. Using this method,

a dose-response curve was first determined for the DOTAP :

DOPE cationic lipid. Dose-response curves were then

examined using endotoxin alone and endotoxin incubated with






81


relatively nontoxic concentrations of lipid (<5 pg/ml).

While the DOTAP : DOPE

1.0



S0.8
E


S0.6


C
c
o 0.4
0


0.2



0.0 -' 1 i. *j i i. I i ii *I 1'3I3
0 .1 1 10 100
DOTAP:DOPE (ng)
Figure 28: Effect of increasing lipid concentration with
endotoxin held constant (0.6 EU) on endotoxin activity,
EU/ml + SEM, n=3, p>0.05 via one way ANOVA for effect of
cationic lipid on endotoxin activity.

cationic lipid was toxic, no significant loss of COS-1 cell

viability was detected at endotoxin concentrations up to

5000 EU/ml in the presence of lipid and no change in

viability was seen with free endotoxin (Figure 29).


Discussion

The interactions of endotoxin with non-viral gene

delivery systems and the resultant decreases in






82


transfection efficiency are of potential concern in the

success of gene delivery. If a fundamental understanding

of the factors that influence endotoxin-delivery system

interaction is gained, vectors might be developed that

limit the impact of endotoxin in these formulations.

120"
A B
1001

80

60

40

20


1 10 100 5 50 500 5000
DOTAP:DOPE (ng/ml) Endotoxin ( gg/ml)

Figure 29: Effect of increasing cationic lipid (panel A)
and endotoxin (panel B) on COS-1 cell survival via MTT
assay. mean + SEM, n=5, p < 0.05 via ANOVA for DOTAP :
DOPE, p > 0.05 for endotoxin via ANOVA.

Fluorescent anisotropy results demonstrate endotoxin

interacts with DOTAP : DOPE cationic liposomes.

Fluorescent anisotropic measurements are well suited for

observing interactions between molecules since the reporter

fluorescent probe is sensitive to its environmental

conditions. Fluorophores have a defined orientation and

preferentially absorb light that is vectored in that






83


orientation. By excitation using polarized light, it is

possible to selectively excite individual molecules. As

molecules undergo faster rotational diffusion in solution,

the time of fluorescence emission and anisotropy decreases.

As the molecular volume increases with interaction, there

is a correlative decrease in rotational diffusion and thus

an increase in the value of the anisotropic measurement.

This anisotropic methodology has been applied to observing

oligonucleotide hybridization in solution (Murakami et al.

1991) and to examine oligonucleotide-dendrimer interaction

(Poxon et al. 1996).

The structure and physical properties of endotoxin

contribute to the interaction with cationic DOTAP : DOPE

liposomes. For instance, the fatty acid chains present on

the lipid A moiety contributes to the amphipathicity of

endotoxin. Endotoxin's amphipathic nature coupled with its

low solubility causes aqueous preparations of endotoxin to

exist in mainly an aggregated state (Takayama et al. 1995).

The prevalence of the aggregated state results in the

clustering of the negative charge. This cluster of charge

should help to contribute to the electrostatic interaction

between the cationic liposomes and the anionic endotoxin.

The LAL assay provided information on the effects of

the endotoxin-cationic lipid interaction on potential






84


biological activity. The interaction of the endotoxin and

cationic DOTAP liposomes did not alter the activity of the

endotoxin as measured by the LAL assay. This implies

either that the interaction does not hinder the toxic lipid

A moiety or that the complex is not stable.

The MTT assay demonstrated the effects of endotoxin-

cationic lipid complex on COS-1 cell viability. Increasing

levels of endotoxin-cationic lipid complex and free

endotoxin had no significant effect upon COS-1 cell

viability. This lack of selective toxicity in an

immortalized cell line has previously been reported,

including HeLa, Vero, 3T3, K562, WI-38, SV1, TX-4, CHO,

P3U3, and R-393 cells (Epstein et al. 1990, ; Cotton et al.

1994). Immortalized cell lines have shown no change in

growth as determined by doubling time, plating density and

confluent density measurements. Rather, the toxic response

occurs in primary cell cultures cells (Epstein et al. 1990;

Cotton et al. 1994). In any case, the effect of endotoxin

in a more complex in vivo system is extremely toxic and

should not be dismissed.

The mechanism of endotoxin-cationic lipid interaction

is thought to be mainly electrostatic, as evidenced by the

trend to decreased interaction after endotoxin

dephosphorylation and ionic strength experiments. However,






85


other forces may be involved due to the inability of high

screening-ion concentrations to completely inhibit the

interaction phenomena and the ability of zwitterionic

liposomes to interact with endotoxin. It should be noted

that there were a different number of particles with a

different exposed surface area at similar weights of

lecithin liposomes and cationic dendrimers. This fact

could change the rate of endotoxin interaction with each

compound. However, the greater change in anisotropy

observed with the cationic dendrimers, which should undergo

solely electrostatic interaction, is most likely indicative

of a greater electrostatic interaction.

Structural flexibility of the vesicular liposome

versus the more static, solid dendrimer may also have an

impact on the interactions. When endotoxin undergoes an

electrostatic interaction with cationic liposomes, this

interaction can result in lower transfection efficiencies,

as measured by enzymatic expression. Furthermore, this

effect may not solely be due to a difference in the

delivery of the complex as supported by these studies.

Previous studies with transiently transfected CHO cells

show that exposure to endotoxin after transfection results

in a stimulation of the transfected product at 10 ng/ml

endotoxin but decreases production at higher levels






86


(Epstein et al. 1990). A trend towards a bimodal response

can be seen in Figure 22, but was not statistically

significant in this study, resulting only in increased

variability.

These results suggest that the effect of endotoxin on

established cell lines is not through a cytotoxic mechanism

but rather a difference in the delivery of the plasmid-

endotoxin-cationic lipid complex. The inclusion of

endotoxin into the cationic liposome complex may alter the

morphological form and overall net charge of the complex,

thereby altering the delivery and subsequent transgene

expression. The displacement of DNA from cationic liposomes

by endotoxin is most likely responsible for the decreased

transfection efficiency in established cell lines, since

lowered delivered plasmid levels would result in decreased

expression.

From the present study it is clear that several

factors besides the plasmid and delivery vector will

influence the activity of non-viral gene delivery systems.

Endotoxin contamination can potentially impact transfection

efficiency via competition with plasmid DNA for cationic

liposome binding, but this would not be expected with

typical GLP or GMP preparations used in clinical studies

where the endotoxin levels range from 10 1000 EU/mg DNA.






87


At these reduced endotoxin levels, the competition between

endotoxin and DNA for the cationic liposome is at best

marginal. An effect can be seen, with increased

transfection variability, at 50 EU/ml. This is a level of

endotoxin contamination that can occur with small scale

plasmid preparations used for in vitro cell transfections,

and potentially affects the results seen in many in vitro

studies.















CHAPTER 5
FOAM FRACTIONATION AS A METHOD TO SEPARATE ENDOTOXIN FROM
RECOMBINANT BIOTECHNOLOGY PRODUCT


Introduction

Biotechnology based therapy, using products isolated

from E. coli, need to be concerned of a potential for

toxicity from the contaminant endotoxin (Weber et al.

1995). Endotoxin, also known as lipopolysaccharide or LPS,

is a gram negative bacterial cell wall component commonly

co-isolated with plasmid DNA and recombinant proteins. It

consists of a polysaccharide component and a covalently

bound lipid component, lipid A. Lipid A is biologically

active and can cause a number of pathophysiological effects

including fever, hypotension, intravascular coagulation and

death (Fletcher, and Ramwell 1980; Aida, and Pabst 1990;

Rietschel et al. 1993; Xing et al. 1994).

The removal of plasmid DNA endotoxin contamination can

be difficult on several accounts (Cotton et al. 1994).

First, the negative charges associated with lipid A will

cause endotoxin to mimic DNA on anion exchange resins.

Second, the large size of endotoxin molecule aggregates



88




Full Text

PAGE 1

\ THE PHARMACEUTICAL STABILITY AND FORMULATION OF PLASMID DNA By SCOTT WILLIAM POXON A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 1999

PAGE 2

This dissertation is dedicated to my wife, Stephanie Poxon, without whose understanding and patience I could not have accomplished this task. As well, I would like to thank my parents and the other students in the Hughes laboratory for their help and support.

PAGE 3

ACKNOWLEDGMENTS I would like to acknowledge my committee members, Dr. Jeffrey Hughes, Dr. Gayle Brazeau, Dr. Edwin Meyer, Dr. Michael Schwartz, and Dr. Ian Tebbett for their critical reading of this manuscript, support and contribution to this project. The Parenteral Drug Association Foundation Biotechnology Grant, the National Institutes of Health (NIH POl-AG 10485), and the University of Florida Gene Therapy Center also financially supported these studies. iii

PAGE 4

TABLE OF CONTENTS page ACKN"OWLEDGMENT S . . . . . . . . . . . . . . . . . . . . iii LIST OF FIGURES .......................................... vi ABSTRACT ................................................. xi 1 INTRO DUCT I ON . . . . . . . . . . . . . . . . . . . . . . 1 Specific Aims and Hypotheses ............................ 1 Background and Significance ............................. 2 Chemical and Physical Stability of DNA .................. 5 Background ........................................... 5 Glycosidic Bond .................................... 7 Phosphodiester Bond .................................. 9 Steric Effects ................................... 10 Oxidative Damage .................................... 10 Lyophilization of DNA ................................. 11 Endotoxin Contamination of DNA ......................... 13 Conclusions ............................................ 15 2 PHYSICAL AND CHEMICAL STABILITY OF PLASMID DNA ........ 16 Background ............................................. 16 Materials and Methods .................................. 21 Plasmid Purification ................................ 21 Liposome Preparation ................................ 22 In vitro Coupled Transcription-Translation .......... 23 Transfection Efficiency Assay ....................... 23 DMED Ass a y . . . . . . . . . . . . . . . . . . . . . 2 5 ~ H Solution Stability Study ......................... 25 ~ emperature Stability Study ......................... 26 Statistical Analysis ................................ 27 Results and Discussion ................................. 27 3 FORMULATION OF PLASMID DNA: THE EFFECT OF LYOPHILIZATION ON PLASMID DNA STABILITY ................ 34 Introduction ........................................... 34 iv

PAGE 5

Materials and Methods .................................. 38 Plasmid Purification ................................ 38 Liposome Preparation ................................ 39 Transfection Efficiency Assay ...................... 40 Differential Scanning Calorimetry Analysis .......... 41 DME D As s a y . . . . . . . . . . . . . . . . . . . . 4 2 Analysis of Hyperchromic Effect ..................... 42 Oxidative Analysis ................................ 43 Circular Dichroism Analysis ......................... 44 Statistical Analysis ................................ 44 Results and Discussion ................................. 44 Conclusion ............................................. 61 4 CHARACTERIZATION OF ENDOTOXIN AND CATIONIC LIPOSOME INTERACTION ........................................... 64 Introduction ........................................... 64 Materials and Methods .................................. 67 Plasmid Purification ................................ 67 Liposome Preparation ................................ 68 Transfection Efficiency Assay ...................... 68 Anisotropy Assay .................................... 70 Endotoxin Dephosphorylation ......................... 71 Endotoxin Assay ..................................... 71 Ce 11 Vi ab i 1 it y Ass a y . . . . . . . . . . . . . . . 7 2 Statistical Analysis ................................ 7 3 Results ................................................ 73 Discussion ............................................. 81 5 FOAM FRACTIONATION AS A METHOD TO SEPARATE ENDOTOXIN FROM RECOMBINANT BIOTECHNOLOGY PRODUCTS ................ 88 Introduction ........................................... 88 Methods ................................................ 91 Plasmid Purification ................................ 91 Foam Fractionation .................................. 92 Plasmid and FITC-Endotoxin Gel Analysis ............. 92 Surface Activity Determination ...................... 93 Particle Size Analysis .............................. 93 Endotoxin Assay ..................................... 93 Protein Analysis .................................... 94 Statistical Analysis ................................ 94 Results and Discussion ................................. 94 Conclusions ............................................. 103 LIST OF REFERENCES ...................................... 107 BIOGRAPHICAL SKETCH ..................................... 115 V

PAGE 6

LIST OF FIGURES Figure 1. Major sites for chemical degradation of DNA. Filled arrows point to potential oxidative damage sites, while open arrows point to potential page hydrolytic damage sites ............................... 6 2. Glycosidic cleavage of cytosine to an aldehyde under alkaline or acidic conditions followed by beta elimination to cleave the phosphate backbone. ............................................ 7 3. Comparison of cotransfection (A) and single plasmid (B) concentration vs activity. Mean SEM, n=4 ............................................. 28 4. Effect of incubation at elevated temperature for 3 weeks on the functional activity of plasmid DNA as measured via coupled transcriptiontranslation. RLUSEM, n=3, 37C is significantly different than 75 and 95C (p<0.01) via Fisher's PLSD ................................................. 29 5. Effect of incubation at various pHs for 3 weeks on the functional activity of plasmid DNA as measured via coupled transcription-translation. RLUSEM, n=6, *=p<0.05 via Fisher's PLSD compared to pH 7 ..................................... 3 0 6. Effect of incubation in pH 3, 2.5 mM citrate buffer on plasmid DNA in a 0.8% agarose gel, stained with ethidium bromide ........................ 31 7. Effect of citrate buffer concentration on pRLCMV plasmid degradation rate at 50C. Mean+ SEM, n=5, r 2 >0. 98 for all fit lines. ...... :-_ ... ..... 32 8. Effect of lyophilization (FD) on plasmid DNA activity, with various amounts lyoprotectant to DNA (w/w). Average SEM. n=5 for all treatments, *=FD DNA significantly lower than Control DNA. p<0.05 via Schefe's multiple comparison T-test .................................... 4 6 vi

PAGE 7

9. Representative thermal analysis scan of salmon sperm DNA ............................................ 49 10. Effect of lyoprotectants on the melting temperature of salmon-sperm DNA. Mean+ SEM, n=3, *=significantly different than unprotected control DNA (p < 0.05) via Fisher's PLSD. 50 11. Effect of lyophilization and subsequent rehydration on the melting of plasmid DNA. Mean SEM, n=3 ............................................. 51 12. Lyophilized plasmid DNA samples, rehydrated in DI water and run an agarose gel. Samples: marker, HindII digested lambda phage marker; FD, lyophilized DNA without DMED treatment; FD DMED, lyophilized DNA with DMED treatment; pH 3, DNA incubated in pH 3.0, 2.5 mM citrate buffer for 15 minutes at 50C with no DMED treatment; pH 3 DMED, DNA incubated in pH 3.0, 2.5 mM citrate buffer for 15 minutes at 50C with DMED treatment before running on gel ................................ 52 13. Effect of lyophilization on plasmid form. Mean SEM, n=3. p > 0. 05 via ANOVA ........................ 53 14. Effect of lyophilization on plasmid DNA absorbance at 260 nm. Average SEM. n=4, FD DNA significantly higher than lyoprotected DNA p< 0 0 5 vi a ANO VA . . . . . . . . . . . . . . . . . . 5 4 15. Wavelength circular dichroism scan of lyophilized plasmid DNA (FD) compared to A-form and B-form plasmid DNA. Each scan is the average of three separate baseline subtracted scans .......... 55 16. Effect of lyoprotection on plasmid DNA conformational change. Panels each signify a separate protectant; A: lactose, B: glucosamine, C: glucose-1-phosphate, and D: glucose. For each panel Protectant:DNA 1, 5, 10, 20 and 40 or 1, 2, and 8 w/w ratios. Cntrl S ~protectant" is protectant:DNA solution (40:1 w/w) non lyophilized. Cntrl FD and Cntrl S ~without protectant" are no protectant, lyophilzed or non lyophilized respectively. n=l0, mean+ SEM .......... 56 vii

PAGE 8

17. Kinetics of plasmid conformational change after rehydration. mean+ SEM, n=3 ........................ 58 18. Effect of rehydration time on plasmid DNA activity. Mean SEM. n=6. No significant difference between O and 24 hours by T-test .......... 59 19. HPLC analysis of hydrolyzed pRL-CMV plasmid DNA. Panel A is a UV analysis. Peaks by retention time: 2.76, solvent front, 3.79 cytosine, 7.85 guanine, 11.15 thymine, 13.00 unidentified, 17.89 adenine. Panel Bis an ECD analysis. Peaks by retention time 2,67 3.21 solvent front, 3.53 5OH cytosine, 3.82 cytosine, 4.78 plasticizer, 7.35 guanine, 7.92 8-OH guanine, 16.99 8-OH adenine and adenine . ................................ 6 O 20: Effect of lyophilization of plasmid DNA on oxidative damage. Positive control was Fe + 3 catalyzed oxidation. Mean SEM, n=5, except control n=3. = p < 0.05 via Fisher's PLSD versus control ....................................... 61 21. Effect of tube type on plasmid DNA bioactivity after lyophilization. Cntrl = non-lyophilized DNA, PP= polypropylene, SP= siliconized polypropylene, PE= polyethylene. Mean SEM, n=5, = p < 0.05 via Fisher's PLSD versus control, X = p < 0.05 versus PP and SP ............... 62 22. Enzyme activity corrected for total cellular protein after transfection of luciferase plasmid (lg) in the presence of endotoxin, with and without DOTAP: DOPE cationic liposomes. with lipid (2 ug/ml), without lipid. RLUSEM, n=4, p<0.05 via one way ANOVA for effect of endotoxin in the presence of cationic lipid .................... 74 23. Effect of increasing DOTAP : DOPE liposomes concentration with FITC-endotoxin held constant (1000 EU) on anisotropy (r). DI Water, 0.5 M NaCl, A 1.0 M NaCl, X 2.0 M NaCl. Inset: Free FITC (22 ng) 1.0 M NaCl. Change in rSEM, n=3, p<0.05 using two way ANOVA for increase in r with increasing lipid under all conditions ................ 76 viii

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24. Effect of increasing luciferase plasmid concentration on anisotropy (r) with constant FITC-endotoxin (1000 EU) and DOTAP: DOPE liposomes (5 g). Change in rSEM, n=3, p<0.05 using two way ANOVA for decrease in r with increasing plasmid ................................... 77 25. Effect of increasing lipid or dendrimer concentration with FITC-endotoxin held constant (1000 EU) on anisotropy (r). Dendrimer (Generation 2), Dendrimer (Generation 4), X lecithin liposomes (0.2 m), A lecithin liposomes (0.8 m), Change in rSEM, n=3, p<0.05 using two way ANOVA for increase in r with increasing concentration under all conditions ................... 78 26. Effect of increasing DOTAP: DOPE liposome concentration with FITC-Endotoxin held constant ( 10 g) .............................................. 7 8 27. Effect of increasing incubation time of alkaline phosphatase and unlabeled endotoxin (33 EU) on anisotropy (r) with constant and NBD labeled DOTAP: DOPE liposome (10 g). Change in r SEM, n=4, p>0.05 via one way ANOVA for decrease in anisotropy over time ................................. 80 28. Effect of increasing lipid concentration with endotoxin held constant (0.6 EU) on endotoxin activity, EU/ml SEM, n=3, p>0.05 via one way ANOVA for effect of cationic lipid on endotoxin activity ............................................. 81 29. Effect of increasing cationic lipid (panel A) and endotoxin (panel B) on COS-1 cell survival via MTT assay. mean SEM, n=5, p < 0.05 via ANOVA for DOTAP: DOPE, p > 0.05 for endotoxin via ANOVA ....................................... 82 30. Effect of foam fractionation on FITC labeled endotoxin levels (initial 100 g/ml n=3) and BSA (initial 910 g/ml, n=3) in a plasmid DNA solution, (25 g/ml), mean SEM. p>0.05 using one way ANOVA for differences in endotoxin concentration over time. p<0.05 for differences in BSA concentration over time ....................... 96 ix

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31. Effect of foam fractionation on endotoxin activity. Mean SEM, p > 0.05 via AAOVA ............ 97 32. Effect of foam fractionation on physical stability of plasmid DNA, molecular weight marker A phage Hind III digest in lane 1; lanes 2 8, fractionation time points 1 g pDNA per lane, in a 0.8% agarose gel stained with ethidium bromide ..... 98 33. Surface tension (dynes/cm) of water, endotoxin, FITC-endotoxin, and BSA (all 1 mg/ml), n=4, mean SEM, = p<0.05 using Fisher's PLSD versus water ................................................ 99 34. Volume weighted particle size of FITC-endotoxin (1 mg/ml) at increasing HCl concentration. Data are shown as meanSEM., p<0.05 via AAOVA, *= p<0.05 using Fisher's PLSD verus 1 M HCl ........... 100 35. Surface tension (dynes/cm) of FITC-endotoxin (1 mg/ml) at various HCl concentrations, n=4, mean SEM, = p<0.05 using Fisher's PLSD versus O m HCl ................................................. 101 X

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy THE PHARMACEUTICAL STABILITY AND FORMULATION OF PLASMID DNA By Scott W. Poxon May, 1999 Chairman: Jeffrey Hughes, Ph.D. Major Department: Pharmaceutics This research examined the stability and formulation of plasmid DNA from a pharmaceutical perspective. Plasmid DNA has the potential to be used as a drug, which could replace missing or damaged proteins that cause disease. As well, plasmid DNA could be used as the basis for new types of vaccines. However, the potential for this new drug can not be realized without research into the formulation and stability of plasmid DNA. These studies examined the xi

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stability of plasmid DNA with respect to environmental factors such as temperature and pH, showing that plasmid DNA is exceedingly stable under many conditions. Furthermore, this research compared the stability of plasmid DNA in both solution and solid lyophilized states. These studies demonstrated that the lyophilization process damages DNA through what appears to be a conformationally induced denaturation, although the lyophilized plasmid DNA is more stable at elevated temperature than plasmid DNA in solution. Finally, it was ascertained that a potential contaminant of plasmid DNA, endotoxin, can decrease DNA transfection efficiency through a competitive electrostatic interaction with a common DNA delivery vector, cationic liposomes. It was also established that the endotoxin contaminant can not be removed from plasmid DNA using foam fractionation. However, the foam fraction method can successfully be used to separate endotoxin from amphiphilic protein products. This research should help to support the future pharmaceutical development of plasmid DNA as a therapeutic modality. xii

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CHAPTER 1 INTRODUCTION Specific Aims and Hypotheses The overall goal of this project was to determine the impact of environmental factors on plasmid DNA stability. Using structural and biofunctional assays, the stability and potential mechanisms of plasmid DNA instability were investigated. This information was then assessed with respect to the pharmaceutical development of plasmid DNA in a therapeutic modality. Four specific hypotheses were investigated by this project. The first hypothesis was that biofunctional assays are more sensitive than structural assays, since damage at one base in the encoding region would not be easily detected by conventional structural methods, but could result in a functionally inactive product. Secondly, it was hypothesized that environmental factors will affect the stability of plasmid in solution. The third hypothesis was that the stability of lyophilized plasmid DNA to environmental factors will be higher compared to plasmid DNA in solution. However, the lyophilization process may 1

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2 damage plasmid DNA via conformational strain caused by the removal of the DNA hydration sphere. Finally, this project tested the hypothesis that increasing levels of endotoxin contamination would result in reduced functionality of plasmid DNA in tissue culture models. To test these hypotheses, the following were the specific aims of this work: 1. To establish and validate a sensitive and specific assay to quantitate the structural and biofunctional stability of plasmid DNA. 2. To determine the optimum storage conditions for plasmid DNA in solution. 3. To determine the influence of the hydration sphere on optimum storage conditions for lyophilized plasmid DNA. 4. To determine the effect of plasmid DNA endotoxin contamination on functionality using specific assays. Background and Significance DNA, or deoxyribonucleic acid, is the genetic material of life. It is used to pass information from one generation to the next, supplying the set of genes needed for the manufacture of further structures in the organism. DNA functions via its capacity to encode a large variety of proteins, where specific sequences of nucleotides in DNA

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3 encode different proteins. Structurally, DNA consists of two antiparallel strands whose sequences are made up of chemically linked subunits, consisting of a nitrogenous base (purine or pyrimidine) attached to a pentose sugar linked together by a phosphate backbone. Each nucleic acid contains one of four types of nitrogenous base: two purines, adenine and guanine, and two pyrimidines, cytosine and thymine. In mammals, chromosomes make up the discrete unit of the genome. Each chromosome consists of a long duplex DNA complexed with protein. Another form of DNA, plasmid DNA, is an autonomous unit that can exist inside a bacterial cell's cytoplasm. These extrachromosomal, double stranded, circular molecules of DNA are self-replicating. During the 1970s, many artificial plasmids were constructed in the laboratory utilizing fragments from naturally occurring plasmids, and are commonly used as vectors for recombinant DNA work and gene therapy. Since DNA is the genetic material, mutations to DNA can potentially have serious consequences. A change in the sequence of DNA causes an alteration of the coded protein, which may cause mutational inactivation. Mutations may either be inherited or induced. In either case, replacement of the mutated DNA via gene therapy can result

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4 in the production of competent protein. This makes gene therapy one of the most exciting and rapidly advancing areas of medicine. Great strides have been recently made in the development of DNA as a therapeutic agent including the successful injection and expression of plasmid DNA into animals (Nabel et al. 1989; Nabel et al. 1990; Nabel et al. 1992; Stewart et al. 1992). These initial successes have quickly led to human clinical trials (Nabel et al. 1993; Caplen et al. 1994). Although there has been some initial investigation into the potential production methods for pharmaceutical grade DNA (Horn et al. 1995; Durland, and Eastman 1998), little information is available concerning plasmid DNA stability in a pharmaceutically acceptable dosage form. For instance, shelf life has not been determined in any potential dosage form, and it is commonly assumed that DNA is stable as either an ethanol precipitate or a frozen buffered solution at neutral to slightly basic pH (Maniatis et al. 1982). However, neither postulate has been thoroughly investigated with respect to the pharmaceutical development of plasmid DNA.

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5 Chemical and Physical Stability of DNA Background The reactivity of the phosphate backbone, the deoxyribose, and the nitrogenous bases that make up the individual components of DNA has been well documented (Shabarova, and Bogdanov 1994). Initial research in this area determined how bases may react with exogenous electrophilic agents, resulting in heterocyclic halogenation or nitration. Other electrophilic reactions include methylation and oxidation. Reactions with amine containing nucleophilic reagents may also result in substitutions under basic conditions (Shabarova, and Bogdanov 1994). Of these reactions, oxidation is most likely of concern in long-term DNA storage since the plasmid isolation process should not result in the addition of exogenous electrophiles (Finnegan et al. 1996). However, plasmid isolation may result in the addition of activated oxygen species through air saturated buffers or the use of organic solvents for extraction resulting in subsequent oxidative damage (Finnegan et al. 1996). Although these previous reactions have generally been characterized experimentally using nucleic acid monomers, it is possible that the reactivity of plasmid DNA may

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6 increase or decrease significantly from the monomers due to steric factors associated with the primary and secondary structures of plasmid DNA. Several specific areas of the DNA have been extensively studied for their lability under physiologic conditions. The sites susceptible to hydrolytic attack and oxidative damage in plasmid DNA can be seen in Figure 1. 3' Figure 1: Major sites for chemical degradation of DNA. Filled arrows point to potential oxidative damage sites, while open arrows point to potential hydrolytic damage sites.

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7 Glycosidic Bond The glycosidic bond is highly stable under neutral or basic conditions, but is extremely sensitive to acid hydrolysis. Acid hydrolysis of the glycosidic bond on nucleotides occurs rapidly with purines (k=104 sec1 ) and more slowly with pyrimidines (k=108 sec1 ) at pH 1.0, 37C, since the purines are better leaving groups than pyrimidines (Lindahl 1993). Under physiologic conditions, it has been predicted that mammalian cells undergo 2,000 to 10,000 cases of hydrolytic depurination followed by repair every day (Lindahl 1993). In pharmaceutical storage of plasmid DNA, however, no repair mechanisms exist, bringing concern about potential degradation of the glycosidic bond. 5 CNHi 6 1 N I 0=~~0~ 0 (>_ ),-1 O=t>-o6 I 3' 5' 6 6I H O=t>-1?-HO H O=t>-o6 J, Figure 2: Glycosidic cleavage of cytosine to an aldehyde under alkaline or acidic conditions followed by beta elimination to cleave the phosphate backbone. After depurination by cleavage of the glycosidic bond, an aldehyde forms at the Cl of deoxyribose, which can then

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8 undergo beta elimination as shown in Figure 2 (McHugh and Knowland 1995). This aldehyde form exists in equilibrium with the cyclic depurinated hemiacetal form with about 1% of the base-free sugar residue in the aldehyde form at any one time Even without chemical catalysis, the weakened DNA chain would undergo the elimination process within a few days rendering the stored plasmid DNA non-functional by blocking DNA polymerase It has been shown that deamination of the bases may also occur in alkaline conditions resulting in potential loss of the encoded protein s functionality This deamination process occurs mainly via acid-catalysis under physiologic conditions (Lindahl 1993) This however, is a slow reaction, with the half life of an individual cytosine residue in single stranded DNA extrapolated to 200 years (Frederico et al 1990) The hydrolytic deamination reaction slows further with double stranded DNA to a half life of about 30 000 years for each cytosine residue (Frederico et al 1990) Deamination occurs at a slower rate than depurination, and is therefore less likely to be the rate-determining step in plasmid degradation during storage

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9 Phosphodiester Bond During prolonged exposure to high temperatures, DNA will progressively melt via first order kinetics followed by heat-induced hydrolysis of the phosphodiester bond with an expected 3,000 fold increase in DNA decay at 100C as compared to 37C (Lindahl 1993). At temperatures above 100C the DNA is very unstable because of both its chemical nature to hydrolyze and the problem of retaining the hydrogen bonding between the two DNA strands at high temperatures. It has also been suggested that increased pressure could help stabilize DNA, since the melting temperature of the double helix is 10C higher at 5,000 atmospheres than at 1 atmosphere (Lindahl 1993). The phosphodiester bond is another potential site for damage in plasmid DNA. It may be broken by beta elimination as well as by acid hydrolysis at a pH below 3. The phosphodiester backbone can also be cleaved by nucleases and by oxidative degradation, but is otherwise as stable at neutral and basic pH as the glycosidic bond and the amine groups on the bases. This explains the reasoning behind current DNA storage paradigms of freezing DNA at neutral to slightly basic pH.

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10 Steric Effects Positive supercoiling can act as a barrier to deamination, depurination and subsequent beta elimination. Protection via supercoiling is due to a change in the plasmid's tertiary structure that results in torsional twisting of the DNA structure. This twisting causes the duplex to cross itself in space and is responsible for the increased pH needed to denature circular DNA as compared to linear DNA. Supercoiling can also protect against thermal DNA degradation (Marguet, and Forterre 1994), suggesting that condensation of plasmid DNA may increase stability during storage. Oxidative Damage Oxidative damage of plasmid DNA may be a problem inherent in long-term storage, particularly if the DNA was exposed to hydroxyl or superoxide radicals (Lindahl 1993). Exposure to activated oxygen radical is ubiquitous inside cells, where DNA repair enzymes exist that reduce the damage to cellular DNA. In long-term storage of plasmid DNA, these repair enzymes will not exist, so oxidative damage can accrue resulting in non-functional protein product. Damage can potentially occur during plasmid DNA isolation with solvents such as phenol and chloroform, and

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11 air-saturated buffers (Lindahl 1993; Finnegan et al. 1996). Exposure to activated oxygen can cause the formation of formamidopyrimidines, purines with opened imidazole rings, that are non-coding residues. As well, hydroxyl radicals can react with guanine, forming 8-hydroxyguanine, which base pairs with adenine rather than cytosine. Therefore, transversion mutations will be generated after replication. Another common oxidative-induced DNA damage is the generation of ring saturated pyrimidines. In this case, losing the 5,6 double bond causes the loss of planar ring structure leaving a non-coding base residue (Lindahl 1993). Lyophilization of DNA Lyophilization of plasmid DNA may be a preferred form of storage, potentially imparting a longer shelf life and greater stability against heat-induced degradation. / Experimental findings support this by suggesting that biological macromolecules demonstrate an increase in stability from the frozen to the lyophilized state (Crowe et al. 1990). The exact mechanism underlying this difference has not been fully elucidated, but most likely relates to the nonfreezable water associated with most biomolecules, including nucleic acids. It is well known that the structure and conformational states of DNA are )

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12 critically dependent on hydration level (Umrania et al. 1995; La Vere et al. 1996). Several forms of DNA secondary structure are known and, under normal physiological conditions, DNA is found in the B-form. This form has a major and a minor groove, with 10 base pairs per turn. The A-form is more compact, with 11 base pairs per turn, and is seen when the presence of 2'hydroxyl groups on the ribose prevents formation of the B-form during complexation with RNA, and under conditions of high salt concentration, or low humidity (<75%), as expected under lyophilized conditions. While the A and B-forms are both right handed helixes, the Z-form is a left handed helix and has the most base pairs per turn of any of the forms, and only one groove. This form has been seen under high salt concentrations and probably does not exist in vivo but could potentially be seen in low concentrations under the high salt conditions seen with lyophilization. The secondary structure of the DNA may affect stability and functionality. Lyophilization causes the removal of the hydration sphere around a molecule. For DNA, it appears that there are approximately 20 water molecules per nucleotide pair bound most tightly to DNA that do not form an ice-like structure upon low temperature cooling. Upon DNA

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13 dehydration at 0% relative humidity only five or six water molecules remain (Falk et al. 1963; Lith et al. 1986; Tao, and Lindsay 1989). Lyophilization may increase the stability of DNA under long-term storage, but may also cause some damage upon the initial lyophilization process, potentially through changes in the DNA secondary structure. Agents that can substitute for nonfreezable water, such as trehalose, can demonstrate cryoprotective properties for DNA and other molecules during lyophilization of intact bacteria (Rudolph et al. 1986; Israeli et al. 1993). Other cryoprotective agents, such as polyols, amino acids, sugars and lyotropic salts are preferentially excluded from contact with protein surfaces but are also capable of stabilizing enzymes during lyophilization by undefined mechanisms (Carpenter and Crowe 1989). It is possible that agents that act as cryoprotectants to proteins may also act to stabilize nucleic acids. Endotoxin Contamination of DNA Since plasmid DNA is typically produced by bacteria, endogenous bacterial products can potentially contaminate the final plasmid DNA product. Typical problem compounds in plasmid DNA preparations include endotoxin, a cell wall component that is pyrogenic in man, and DNase, a ubiquitous

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14 bacterial enzyme that can degrade the final plasmid product. Endotoxin from the cell wall of Gram-negative bacteria, such as the E. coli typically used to prepare plasmid DNA, may be the most serious concern for gene therapy because of its extreme toxicity. Several plasmid DNA preparation methods have been examined for level of endotoxin contamination (Weber et al. 1995). Endotoxin has a relatively low lethal dose for 50% of tested animals (LD 50 ) in rats of 3 mg per kg (Shibayama et al. 1991) and an LD 50 in dogs of only 1 mg per kg (Fletcher, and Ramwell 1980). The endotoxin, or lipopolysaccharide, of E. coli consists of a polysaccharide component and a covalently bound lipid component termed lipid A. The lipid A portion on the endotoxin is biologically active and can cause a number of pathophysiological effects including fever, hypotension, intravascular coagulation and death (Fletcher, and Ramwell 1980; Aida, and Pabst 1990; Rietschel et al. 1993; Xing et al. 1994). The endotoxins are acute inflammatory mediators, elevating levels of cytokines (Xing et al. 1994). Furthermore, an increasing level of endotoxin contamination has been shown to decrease transfection efficiency using DOTAP: DOPE (dioleoyl glycero trimethylammonium propane : dioleoyl glycero phosphoethanolamine) cationic liposomes to deliver plasmid

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15 DNA (Weber et al. 1995). Besides having an easy way to detect endotoxin, there are several ways to remove endotoxin from DNA samples. Phase separation using Triton X-114 has been used to reduce endotoxin levels for both protein and plasmid DNA (Aida, and Pabst 1990; Manthorpe et al. 1993). The use of these methods to reduce endotoxin contamination may be of importance in the potential usefulness of plasmid DNA. Conclusions It is important to understand the effects of the formulation parameters such as pH, temperature, and buffer composition on plasmid DNA stability. Furthermore, the differences between solution and lyophilized plasmid stability must be taken into account to determine the optimal plasmid DNA storage conditions. This research will address the stability of plasmid DNA by examining plasmid structural integrity and biofunctionality. This distinction is important, since current structural methods will not always detect changes in the functionality of the final protein product that can be produced by damage to the plasmid DNA. Using the information provided by these studies, it should be possible to develop a rational basis for the plasmid DNA manufacturing process.

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CHAPTER 2 PHYSICAL AND CHEMICAL STABILITY OF PLASMID DNA Background Before the benefits of gene therapy can be realized on a large scale, the pharmaceutical community needs to ascertain the optimal storage conditions for plasmid DNA so as to prolong shelf life to the greatest possible extent. While a great deal of work has gone into the study of individual nucleotides, little has been accomplished to date regarding the bioactivity of plasmid DNA, or the ability of plasmid DNA to code for a functional protein, as a marker for shelf life. Other methods have been used to examine DNA stability, including agarose gel electrophoresis, HPLC, Southern blotting, and PCR (Niven, Pearlman et. al 1998; Strege and Lagu 1991; Thierry, Lunardi-Iskandar et. al 1995). However, a bioactivity based method will detect changes not only in physical stability of plasmid DNA but also will detect cellular changes in the DNA handling, including, but not limited to, transport, transcription, and translation. 16

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17 The stability of the subunits of plasmid DNA, the individual purine and pyrimidine bases, and their attachment to the ribose sugar and phosphate backbone are fairly well understood. The glycosidic bond is highly stable under neutral or basic conditions, but is extremely sensitive to acid hydrolysis. Acid hydrolysis of the glycosidic bond on nucleotides occurs rapidly with purines ( k=l04 sec1 ) and more slowly with pyrimidines ( k=l08 sec1 ) at pH 1.0, 37C, since the purines are better leaving groups than pyrimidines (Lindahl 1993). Under physiologic conditions, it has been predicted that mammalian cells undergo 2,000 to 10,000 cases of hydrolytic depurination followed by repair every day (Lindahl 1993). In pharmaceutical storage of plasmid DNA, however, no repair mechanisms exist, bringing concern about potential degradation of the glycosidic bond. After depurination by cleavage of the glycosidic bond, an aldehyde forms at the Cl of deoxyribose, which can then undergo beta elimination as shown in Figure 2 on page 7 (McHugh, and Knowland 1995). This aldehyde form exists in equilibrium with the cyclic depurinated form, with about 1% of the base-free sugar residue in the aldehyde form at any one time. Even without chemical catalysis, the weakened DNA chain will undergo the elimination process within a few

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18 days. This would render the stored plasmid DNA non functional by blocking DNA polymerase. It has been shown that deamination of the bases may occur in alkaline conditions, resulting in potential loss of the encoded protein's functionality. This process can also occur via acid hydrolysis under physiologic conditions, where the half-life of an individual cytosine residue in single stranded DNA has been extrapolated to 200 years (Frederico et al. 1990). This hydrolytic deamination reaction slows further with double stranded DNA to a half life of about 30,000 years for each cytosine residue (Frederico et al. 1990). This reaction occurs at a slower rate than depurination and is therefore less likely to be the rate-determining step in plasmid degradation during storage. Depurination rates may be effected by changes in the formulation of plasmid DNA. During prolonged exposure to high temperatures, DNA will progressively melt via first-order kinetics followed by heat-induced hydrolysis of the phosphodiester bond, with an expected 3,000 fold increase in DNA decay at 100C as compared with 37C (Lindahl 1993) This rate increase essentially follows the 10C rule with a 3-fold rate increase per 10C temperature increase. At temperatures above 100C the DNA is very unstable because of both its

PAGE 31

19 chemical nature to hydrolyze and the problem of retaining the hydrogen bonding between the two DNA strands at high temperatures. It has also been suggested that increased pressure could help stabilize the DNA, since the melting temperature of the double helix is 10C higher at 5,000 atmospheres than at 1 atmosphere (Lindahl 1993). The phosphodiester bond is another potential damage site for plasmid DNA. It may be broken by beta elimination as well as by acid hydrolysis at a pH below 3. The phosphodiester backbone can also be cleaved by nucleases and by oxidative degradation, but is otherwise as stable at neutral and basic pH as the glycosidic bond and the amine groups on the bases. This explains the reasoning behind current DNA storage paradigm of frozen DNA at neutral to slightly basic pH. Positive supercoiling can act as a barrier to deamination, depurination and subsequent beta elimination. This supercoiling protection is due to a change in the plasmid's tertiary structure that results in torsional twisting of the DNA structure. This twisting causes the duplex to cross itself in space and this conformationally induced steric protection is responsible for the increased pH needed to denature circular DNA as compared to linear DNA. Supercoiling can also protect against thermal DNA

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20 degradation (Marguet, and Forterre 1994). If supercoiling exerts a protective effect, then it follows that DNA delivery systems that further condense supercoiled plasmid DNA may additionally increase stability during storage. As well, supercoiled plasmid DNA may consequently be more than the monomers that were originally used to determine DNA stability. While it is possible to examine plasmid DNA stability through examination of any of the above individual degradation pathways, a better way to examine DNA stability may be through the examination of plasmid DNA bioactivity, by assaying for enzymatic activity of the plasmid's encoded gene. Assaying for activity of the plasmid DNA will allow a comprehensive determination of degradation that will include all types of damage that affect the expressed gene rather than just the several specific types of damage previously mentioned. The activity of the plasmid can be determined through one of several bioactivity assays. First, plasmid DNA can be assayed using an in vitro coupled transcription-translation system. The in vitro system allows a one tube determination of activity that is not dependent on any particular type of cell system for expression. Rather, the in vitro system includes all the necessary enzymes and cofactors to transcribe the plasmid

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21 DNA to mRNA and to further translate the mRNA to protein, which can then be assayed. This method's main disadvantage is due to variability induced by the ~stop-time" of the reaction. If the reaction is stopped at different times, the resulting enzymatic activity will be effected. The second method for determining the bioactivity of the plasmid is using a mammalian cell based tissue culture transfection assay. This method requires that the plasmid DNA be complexed with a delivery agent before it is transducted into a mammalian cell line. The cell line will then express the protein of interest, which can then be assayed. Several main types of DNA damage are expected, as discussed previously. This study will examine the stability of plasmid DNA using both types of activity assays. Materials and Methods Plasmid Purification Vectors pGL3 plasmid, pSPluc+, and pRL-CMV (Promega, Madison, WI), respectively encoding for photinus luciferase, photinus luciferase and renilla luciferase, were grown in E. coli DH5a cells (Promega, Madison, WI). The transformed bacteria were cultivated in Lura Bertina (LB) broth containing 100 g/ml ampicillin to select for~

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22 lactamase encoding plasmid. The plasmids were isolated via an alkaline lysis method and purified using a silica slurry column {Wizard Plus Megapreps, Promega, Madison, WI). The plasmid was stored in TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 7. 4) Liposome Preparation Lipids were obtained from Avanti Polar Lipids {Alabaster, AL). Cationic DOTAP: DOPE {dioleoyl glycero trimethylammonium propane: dioleoyl glycero phosphoethanolamine) liposomes were prepared using the hand-shaking method {New 1990). DOTAP: DOPE (10 mg DOTAP, 10 mg DOPE) was added to 10 ml of chloroform and introduced into a 250 ml round bottom flask. The chloroform was evaporated using a rotary evaporator at 60C. The lipid film was hydrated with 10 ml of distilled water and shaken for 30 minutes at 60C. Lipids were sized by extrusion, six times through 600 nm polycarbonate filters {Poretics Corp, Livermore, CA). Sizes were confirmed with a laser light scattering particle sizer using volume-weighted distribution {Nicomp 380ZLS, Santa Barbara, CA).

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23 In vitro Coupled Transcription-Translation The coupled transcription-translation assay was carried out using a commercially available kit (Promega, Madison, WI). Essentially, pGL3 plasmid DNA (100 g) was incubated for 60 minutes at 30C with a proprietary mixture containing wheat germ extract, RNA polymerase, amino acids, and a ribonuclease inhibitor. The resulting protein product, luciferase, was then assayed for functionality using a luminescence spectrophotometer (Monolight 2010, Analytical Luminescence Laboratory, San Diego, CA). The light output was measured for 10 seconds and the results integrated to yield the activity. Transfection Efficiency Assay Untreated pGL3 (0.05 g/ml) and treated pRL-CMV (0.5 g/ml) were combined with cationic lipid mixture (DOTAP DOPE 1:1, 2 g/ml each) and incubated for 15 minutes to allow interaction of the anionic plasmid with the cationic liposomes. The 10:1 concentration ratio was as suggested by the manufacturer to achieve a linear relationship between treated renilla luciferase plasmid concentration and renilla luciferase / photinus luciferase activity ratio by decreasing crosstalk between the promoters. These mixtures were then co-transfected into COS-1 cells (ATCC,

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24 Rockville, MD) that had been plated 24 hours earlier at 3x10 4 cells/ml in 24 well tissue-culture dishes. All transfections were effected using serum free DMEM (Dulbecco's Modified Eagle's Medium) supplemented with penicillin G (100 units/ml) and streptomycin (100 g/ml ) The cells were subsequently incubated for 5 hours in a humidified, 5% CO 2 incubator at 37C. The media was aspirated and the cells washed with PBS (phosphate buffered saline) before being replaced by DMEM supplemented with 10% FBS (Fetal Bovine Serum), penicillin G (100 units/ml) and streptomycin (100 g/ml). The cells were incubated for an additional 19 hours, washed with 1 ml PBS, and then lysed with Passive Lysis buffer (200 L, Promega, Madison, WI). Photinus luciferase and renilla luciferase activities were determined by analyzing 5 L of the lysate using the Dual Luciferase Assay kit (Promega, Madison, WI) and a luminescence spectrophotometer (Monolight 2010, Analytical Luminescence Laboratory, San Diego, CA). The light output was measured for 10 seconds and the results integrated to yield the activity. All pRL-CMV, renilla luciferase plasmid, measurements were standardized to the ratio of renilla luciferase (rluc treated) activity to photinus luciferase (luc control) activity. Essentially, the ratio

PAGE 37

25 of rluc/luc is linear to the concentration of rluc added to COS-1 cells. Therefore, a linear regression of rluc/luc ratio at various concentrations of rluc and inactivated rluc (cut with restriction enzymes), with luc held constant, allows a determination of the apparent concentration of the treated pRL-CMV plasmid. DMED Assay Treated pGL3 DNA was incubated with N,N' dimethylethylenediamine (100 rnM, pH 7.4, 30 minutes at 37C), cleaving abasic sites via P-elimination (McHugh, and Knowland 1995). The resultant DNA was then visualized using a 0.8% agarose gel. The gel was electrophoresed at 1 V/cm for 16 hours to affect good separation between the supercoiled, relaxed circular and linear forms of the plasmid DNA. The gel was then stained with ethidium bromide and photographed on an UV light box using a DC40 digital camera (Kodak, Rochester, NY). pH Solution Stability Study The effect of pH upon plasmid DNA bioactivity was studied using a multiple component buffer (Schrier et al. 1993). Poly-B buffer, equimolar sodium citrate, sodium succinate, Tris, HEPES, imidazole, histidine, and glycine at various pHs, was added to pSP6-luc plasmid DNA to a

PAGE 38

26 final concentration of 10 rnM each salt and 0.25 mg/ml plasmid DNA. This mixture was incubated at room temperature for three weeks, with the assumption that pH did not change over time. The plasmid was then precipitated using 70% ethanol, and the pellet rehydrated in TE buffer (10 rnM Tris-HCl, 1 rnM EDTA, pH 7.4) before analysis by the coupled transcription-translation method. The potential for buffer catalysis of plasmid DNA was examined using a citrate buffer at four different concentrations, all at pH 3, 50C. Plasmid, pRL-CMV, was incubated with the various buffers, with aliquots withdrawn over a course of five hours. The aliquots were returned to neutral pH by the addition of excess TE buffer, pH 7.4. The pH was verified to be greater than 7.0 using litmus paper. These samples then were assayed using the cotransfection method. Temperature Stability Study The effect of temperature upon plasmid DNA bioactivity was examined after a three-week incubation at four temperatures: 25, 37, 75 and 95C. Plasmid samples, pSP6luc, were stored in the humidified incubator at 1 mg/ml in TE buffer in sealed 1 ml cryogenic screw-top vials. After

PAGE 39

27 the three-week period, all samples were cooled and assayed using the coupled transcription-translation method. Statistical Analysis Statistical analysis between the various treatments was conducted using analysis of variance and Fisher's protected least significant difference (PLSD) post-hoc tests where appropriate (StatView v4.5, Abacus Concepts, Berkley, CA), with p<0.05 considered statistically significant. Results and Discussion The luciferase cotransfection method used two plasmids coding for two different enzymes (fixed concentration photinus and varying concentration renilla luciferase) to help control for variability by comparing the ratio of the two enzymes activity after transfection. The raw data was analyzed by comparing only the activity of the renilla luciferase. The luciferase cotransfection method of normalizing transfection efficiency was compared to single transfection, raw, non-normalized data shown in Figure 3. It can be seen that the normalized cotransfection data shows a better fit and shows less variability than the single plasmid raw data. The percent coefficient of variation (CV) and percent bias seen with both methods at

PAGE 40

28 each measured concentration can be seen in Table 1. The trend toward lower CV at plasmid concentrations above 0.13 g / ml with the cotranfection suggests that the luciferase cotransfection method is more sensitive than the single plasmid method Another trend is systematic bias below the fit line suggesting that the normalized method is more reproducible than the single plasmid luciferase methods As -,-----------------, B 10 -,---------------~ R 2 = 0 9982 3 2 8 ;! 6 4 2 R 2 = 0 9575 0 0 0 1 0 2 0 3 0 4 0 5 0 0 0 1 0 2 0 3 0 4 pRL-CIIV (ug) p R L-CIIV (ug) Figure 3: Comparison of cotransfection (A) and single plasmid (B) concentration vs activity. Mean+ SEM, n=4 Table 1: Comparison of variance and bias between the luciferase cotransfection and single plasmid methods % CV % Bias g DNA C otransfection Singl.e Cotransfect i on S i ngl.e 0 03 42 27 21 8 0 06 57 40 20 25 0.13 21 29 4 26 0.25 21 20 3 27 0 50 4 38 0 9 Extended incubation of pSP6 luc plasmid DNA for three weeks at elevated temperature can result in a significant decrease in the bioactivity of the treated plasmid at 0 5

PAGE 41

29 elevated temperatures (Figure 4) This degradation would be expected through heat-induced hydrolysis of the phosphodiester bond The bioactivity assay demonstrated a three order of magnitude decrease in plasmid DNA activity after incubation at 95C, which is in line with an expected 3,000 fold increase in DNA decay at 100C compared to 37C as previously reported in the literature (Lindahl 1993). 1 Q6 10 5 :::> ...J 10 4 a::: 1 Q 3 10 0 25 37 75 95 Storage Temperature (C ) Figure 4: Effect of incubation at elevated temperature for 3 weeks on the functional activity of plasmid DNA as measured via coupled transcription-translation. RLU~SEM, n=3, 37C is significantly different than 75 and 95C (p<0.05) via Fisher's PLSD The effect of pH on pSP6-luc plasmid DNA bioactivity was also examined (Figure 5). It can be seen that the

PAGE 42

30 plasmid DNA remained exceedingly stable across a wide pH range. Degradation was seen only at pH 1 and 2. This is not suprising, since the glycosidic bond is reported in the literature to be highly stable under neutral or basic conditions, but is reported to be extremely sensitive to acid hydrolysis (Lindahl 1993). Furthermore, the phosphodiester bond may be broken by beta elimination as well as by acid hydrolysis at a pH below 3. The loss of bioactivity seen at low pH is most likely due to a combination of the two degradation pathways. 10 6 10 5 10 4 :::, _J 10 3 0::: 10 2 10 1 0 1 2 4 6 7 8 10 Storage pH Figure 5: Effect of incubation at various pHs for 3 weeks on the functional activity of plasmid DNA as measured via coupled transcription-translation. RLU+SEM, n=6, *=p<0.05 via Fisher's PLSD compared -to pH 7

PAGE 43

31 Using the DMED assay to examine the mechanism of acidic plasmid DNA degradation, it can be seen that nearly 100% of the plasmid DNA is depurinated at one or more sites per plasmid molecule after 15 minutes (Figure 6). This degradation rate is reasonable compared to published rates with a T 1 ; 2 of 16 minutes at pH 1.0, 37C for depurination (Lindahl 1993). Standard 0 15 30 60 120 240 Minutes oc NL SC DMED 0 15 30 60 120 240 Minutes Figure 6: Effect of incubation in pH 3, 2.5 rnM citrate buffer on plasmid DNA in a 0.8% agarose gel, stained with ethidium bromide. It was also interesting to note that besides undergoing a pH dependent degradation, plasmid DNA may also be subject to buffer catalysis. Statistically different degradation rates were seen when incubating plasmid DNA,

PAGE 44

32 pRL-CMV, in a 50C, pH 3.0 solution, buffered with various amounts of citrate (Figure 7). Degradation rates appear to be first-order exponential and increase with citrate buffer concentration through an undetermined mechanism. 3 -.----------------------, 2.5 > :::E 2 _J a::: B 1.5 1 0.5 -+-----.------.----,-----,----r-------1 0 5 10 15 20 25 30 Incubation Time (min.) mM Figure 7: Effect of citrate buffer concentration on pRL-CMV plasmid degradation rate at 50C. Mean+ SEM, n=5, r 2 >0.98 for all fit lines. In conclusion, plasmid DNA is exceedingly stable in solution over a wide range of pH and temperature extremes. However, the choice of buffer is of more importance than has typically been considered in the past. Based on the literature review and the data presented here, it is most advisable to formulate plasmid DNA at neutral to slightly basic conditions, using previously tested buffers. With buffers that exhibit protective effects, a high concentration of buffer would be preferable. With buffers that that exhibit buffer catalysis, a low buffer

PAGE 45

33 concentration that will minimize any potential buffer effects should be used.

PAGE 46

CHAPTER 3 FORMULATION OF PLASMID DNA: THE EFFECT OF LYOPHILIZATION ON PLASMID DNA STABILITY Introduction Gene therapy is one of the fastest growing areas in therapeutics. While rapid progress has occurred in this field, the pharmaceutical community has not addressed all concerns in detail. Of particular interest is the final dosage form. It is well understood that naked DNA introduced into a patient's circulatory system does not reach enough of the appropriate cells and therefore has little chance of affecting most disease processes (Friedmann 1997). This has led to the development of a number of gene delivery vectors, such as cationic liposomes. Liposomes are considered one of the more promising systems for use in gene delivery (Xu, and Szoka 1996). Unfortunately, cationic liposomes complexed with plasmid DNA are not stable for long-term storage, undergoing aggregation over time (Anchordoquy et al. 1997). There have been several studies with non-ionic liposomes, which may apply to the stability of the 34

PAGE 47

35 DNA/cationic liposome mixture during the lyophilization process. Lyophilization of non-ionic liposomes in the presence of carbohydrates has been cited as one of the most promising ways to keep the liposome stable under long term storage (Williams, and Polli 1984). The lyophilization process, without lyoprotectants, can lead to the leakage of the inner aqueous phase due to liposomal fusion and phase separation of liposomal membranes during drying and rehydration (Crowe et al. 1988). Using carbohydrates as a lyoprotectant will prevent mechanical rupture of the liposomal membrane, caused by ice crystals, during the freezing process and will prevent membrane disruption during drying and rehydration by maintaining the membrane in a flexible state (Ozaki, and Hayashi 1997). Although DNA is complexed with cationic liposomes, not entrapped, it has been shown that the liposome/DNA complex requires lyoprotectants to maintain transfection efficiency after lyophilization (Anchordoquy et al. 1997). Moreover, other gene delivery systems containing polyethylenimine, polylysine, and adenovirus particles require cryoprotection to maintain transfection efficiency (Talsma et al. 1997 ) However, it is unclear if this effect is due only to stabilization of the liposomal component of the mixture or if the plasmid DNA component may itself be damaged by

PAGE 48

36 lyophilization in the absence of carbohydrate lyoprotectants. Lyophilized plasmid DNA may be a preferred form of storage, potentially imparting a longer shelf life and greater stability against heat-induced degradation, since biological macromolecules demonstrate an increase in stability from the frozen to the lyophilized state (Crowe et al. 1990). The exact mechanism underlying this difference has not been fully elucidated, but most likely relates to the nonfreezable water associated with biomolecules, including nucleic acids. It is well known that the structure and conformational states of DNA are critically dependent on hydration level (Umrania et al. 1995; La Vere et al. 1996). Several forms of DNA secondary structure are known and under normal physiological conditions most DNA is found in the B-form. The B-form has a major and a minor groove with 10 base pairs per turn. The A-form is more compact with 11 base pairs per turn. It is observed when DNA is complexed with RNA, due to the presence of 2'hydroxyl groups on the ribose preventing formation of the B-form during complexation. The A-form is also seen under conditions of low humidity (<75%) or high salt concentration (Umrania et al. 1995; La Vere et al. 1996). This A-form secondary

PAGE 49

37 structure is therefore observed under lyophilized conditions. While the A and B-forms are both right handed helixes, the Z-form is a left handed helix and has the most base pairs per turn of any of the forms and only one groove. This form has been seen under high salt concentrations and probably does not form in vivo but could potentially be seen in low concentrations under the high salt conditions seen with lyophilization. The secondary structure of the DNA may effect stability and functionality. Lyophilization causes the removal of the hydration sphere around a molecule. For DNA, it appears that there are approximately 20 water molecules per nucleotide pair bound most tightly to DNA that do not form an ice-like structure upon low temperature cooling. Upon DNA dehydration at 0% relative humidity only five or six water molecules remain (Falk et al. 1963; Lith et al. 1986; Tao, and Lindsay 1989). Lyophilization may increase the stability of DNA under long-term storage because of decreased water activity, but may also cause some damage upon the initial lyophilization process, potentially through changes in the DNA secondary structure. Agents that can substitute for nonfreezable water, such as trehalose, can demonstrate cryoprotective properties for

PAGE 50

38 DNA and other molecules during lyophilization of intact bacteria (Rudolph et al. 1986; Israeli et al. 1993). Other cryoprotective agents, such as polyols, amino acids, sugars and lyotropic salts are preferentially excluded from contact with protein surfaces but are also capable of stabilizing enzymes during lyophilization by undefined mechanisms (Carpenter, and Crowe 1989). It is possible that agents that act as cryoprotectants to proteins may also act to stabilize nucleic acids. Materials and Methods Plasmid Purification Both pGL3 plasmid and pRL-CMV (Promega, Madison, WI), respectively encoding for photinus luciferase and renilla luciferase, were grown in E. coli JM109 cells (Promega, Madison, WI). The transformed bacteria were cultivated in Lura Bertina (LB) broth containing 100 g/ml ampicillin to select for P-lactamase encoding plasmid. The plasmids were isolated via an alkaline lysis method and purified using a silica slurry column (Wizard Plus Megapreps, Promega, Madison, WI). The plasmid was stored in TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 7.4). Plasmid, pRL-CMV, that was to be lyophilized was aliquoted (50 l) into 1.5 ml

PAGE 51

39 polypropylene tubes (Sarstadt) and frozen at -80C. The frozen samples were then lyophilized for 24 hours under vacuum using a Savant Centrivap and a -105C cold trap. Lyophilized samples were rehydrated with distilled water immediately before use. Liposome Preparation Lipids were obtained from Avanti Polar Lipids (Alabaster, AL). Cationic DOTAP : DOPE (dioleoyl glycero trimethylammonium propane : dioleoyl glycero phosphoethanolamine) liposomes were prepared using the hand-shaking method (New 1990). DOTAP: DOPE (10 mg DOTAP, 10 mg DOPE) was added to 10 ml of chloroform and introduced into a 250 ml round bottom flask. The chloroform was evaporated using a rotary evaporator at 60C. The lipid film was hydrated with 10 ml of distilled water and shaken for 30 minutes at 60C. Lipids were sized by extrusion 6 times through 600 nm polycarbonate filters (Poretics Corp, Livermore, CA). Sizes were confirmed with a laser light scattering particle sizer using volume-weighted distribution (Nicomp 380ZLS, Santa Barbara, CA).

PAGE 52

40 Transfection Efficiency Assay Untreated pGL3 (0.05 g/ml) and treated pRL-CMV (0.5 g/ml) were combined with cationic lipid mixture (DOTAP DOPE 1:1, 2 g/ml each) and incubated for 15 minutes to allow interaction of the anionic plasmid with the cationic liposomes in TE buffer (Tris HCl, 10 mM; EDTA, 1 mM, pH 7.4). The mixtures were then co-transfected into COS-1 cells (ATCC, Rockville, MD) that had been plated 24 hours earlier at 3x10 4 cells/ml in 24 well tissue-culture dishes. All transfections were effected using serum free DMEM (Dulbecco's Modified Eagle's Medium) supplemented with penicillin G (100 units/ml) and streptomycin (100 g/ml). The cells were subsequently incubated for 5 hours in a humidified, 5% CO 2 incubator at 37C. The media was aspirated and the cells washed with PBS (phosphate buffered saline) before being replaced by DMEM supplemented with 10% FBS (Fetal Bovine Serum), penicillin G (100 units/ml) and streptomycin (100 g/ml). The cells were incubated for an additional 19 hours, washed with 1 ml PBS, and then lysed with Passive Lysis buffer (200 L, Promega, Madison, WI). Photinus luciferase and renilla luciferase activities were determined by analyzing the 5 L of the lysate using the Dual Luciferase Assay kit (Promega, Madison, WI) and a

PAGE 53

41 luminescence spectrophotometer (Monolight 2010, Analytical Luminescence Laboratory, San Diego, CA). The light output was measured for 10 seconds and the results integrated to yield the activity. All pRL-CMV, renilla luciferase plasmid, measurements were standardized to the ratio of renilla luciferase (rluc treated) activity to photinus luciferase (luc control) activity. Essentially, the ratio of rluc/luc is linear to the concentration of rluc added to COS-1 cells. Therefore, a linear regression of rluc/luc ratio at various concentrations of rluc and inactivated rluc (cut with restriction enzymes), with luc held constant, allows a determination of the apparent concentration of the treated pRL-CMV plasmid. Differential Scanning Calorimetry Analysis Salmon sperm DNA (Sigma Chemical Company, St. Louis, MO), was lyophilized in the presence of lyoprotectant (1:1 w/w). Aliquots of 12-15 mg DNA were then placed in crimped sample pans and the melting profiles determined using a Seiko 220C Differential Scanning Calorimeter (Tokyo, Japan). Scans were carried out over a -10C to 150C temperature range at a heating rate of 5 per minute. Nitrogen was used as a purge gas at a flow rate of 100 ml/min. Only a single scan could be accomplished per

PAGE 54

42 sample, due to thermal degradation of the DNA/carbohydrate mixtures. Onset of melt was determined using the manufacturers software. DMED Assay Treated pRL-CMV DNA was incubated with N,N' dimethylethylenediamine (100 rnM, pH 7.4, 30 minutes at 37C), cleaving abasic sites via P-elimination (McHugh, and Knowland 1995). The resultant DNA was then visualized using a 0.8% agarose gel in TBE buffer (50 rnM Tris, 50 rnM borate, 1 rnM EDTA, pH 7.8). The gel was electrophoresed at 1 V/cm for 16 hours to affect good separation between the supercoiled, relaxed circular and linear forms of the plasmid DNA. The gel was then stained with ethidium bromide (0.5 g/ml) and photographed on a UV light box using a DC40 digital camera (Kodak, Rochester, NY). Analysis of Hyperchromic Effect Hyperchromic effect was examined using a UV/Vis spectrophotometer at 260 nm (Lambda 3, Perkin Elmer). Analysis consisted of comparing the change in absorbance at 260 nm between lyophilized and non-lyophilized samples at the same 1 mg/ml concentration of plasmid DNA.

PAGE 55

43 Oxidative Analysis Oxidative analysis of plasmid DNA was determined using an isocratic HPLC method. Plasmid DNA, 100 g, was first chemically hydrolyzed in 1 ml of 60% formic acid at 150C under vacuum for 45 minutes. Samples were then lyophilized to remove all formic acid and rehydrated in 2 ml of mobile phase (50 rnM sodium acetate, 1 rnM EDTA, 2% methanol, pH 5.5). The samples were then analyzed by HPLC. The HPLC conditions consisted of a 50 l loop, a 0.8 ml/min flow rate, and a 27C column temperature. The column used was a reverse phase C18 column, 25 cm, 5 m (Microsorb MV, Rainin, Woburn MA), with a C18 guard column. Detection was accomplished using a BAS UV-8 detector (West Lafayette, IN) at 254 nm for non-oxidized bases and a BAS LC-4B amperometric detector with a range of 10 nA at 750 mV, using a Ag/AgCl electrode with a glassy carbon working electrode, for oxidized bases. Control oxidized DNA was prepared using a Fenton type iron-catalyzed reaction. Ferric chloride (1 m) and hydrogen peroxide (3%) were incubated with pRL-CMV plasmid (final concentration 1 g/l) on ice for 15 minutes. The plasmid was then purified by ultrafiltration using a 1000 MWCO spin filter (Amicon).

PAGE 56

44 Circular Dichroism Analysis Circular dichroism was analyzed using a Jacso J-500C spectrapolarimeter (Easton, MD). Scans were performed using a 1 nm bandwidth, automatic slit-width, 50 mdeg/fs sensitivity, at 20 nm/min, over a 300-250 nm wavelength range. All scans were baseline subtracted against DI water and each data point was reported as the average of at least two scans. Plasmid DNA conformational change was observed at 270 nm. Samples were analyzed at 0.5 mg/ml pRL-CMV plasmid DNA in a 1 cm cuvette, comparing various weight/weight ratios of pRL-CMV plasmid DNA to lyoprotectant. Lyophilized samples were rehydrated immediately before analysis. Statistical Analysis Statistical analysis between the various treatments was conducted using analysis of variance and Fisher's protected least significant difference (PLSD) post-hoc tests where appropriate (StatView v4.5, Abacus Concepts, Berkley, CA ) with p<0.05 considered statistically significant. Results and Discussion A biofunctional assay was devised to detect damage due to lyophilization of plasmid DNA. This method is a

PAGE 57

45 variation on the conventional cell transfection method, using two plasmids rather than one. By cotransfecting both control and treated plasmid the variation in transfection efficiencies can be taken into account. The treatment plasmid was the pRL-CMV construct (Promega, Madison, WI), containing the renilla luciferase gene driven by a cytomegalovirus promoter. This version of luciferase was isolated from the sea pansy, a species of bioluminescent coral. The control plasmid was the pGL3 construct (Promega). The construct encodes the photinus luciferase gene driven by a simian virus promoter. The two versions of luciferase each require different cofactors and can be assayed in the same test tube, since the conditions appropriate for the renilla luciferase reaction quench the photinus luciferase reaction. The two plasmid constructs were transfected into COSl cells, an African green monkey kidney cell line, that expresses the large T antigen transcription factor needed to drive the SV40 promoter. This transcription factor was capable of upregulating the CMV promoter, as well (Soneoka et al. 1995). The transfections were liposome mediated with the optimal ratio of lipid to pRL-CMV to pGL3 for detection as suggested by the manufacturer. This ratio was empirically based upon the linearity of concentration to activity standard curves

PAGE 58

46 (Figure 3) and the need to prevent cross talk between the different promoters present in the two vectors. The cells were transfected in serum free media and were incubated for 0 7 _0.6 0 5 CD .. 0.4 0.3 0 0 2 0.1 0 Cntr1 4 2 0 1 Glucose:DNA (wlw) 0 7 0 6 10 5 .. 0 4 c( 0.3 z 0 0 2 0 1 0 Cntr1 4 2 0.1 Glucosamlne : DNA (w/w) 0 1 ~ ------------~ 0 6 10 5 .. 0 4 0 3 0 0 2 0 1 0 Cntr1 4 2 Lactose : DNA (wlw) 0 1 0 1 ~-----------------, 0 6 10 5 .. 0 4 0 3 0 0 2 0 1 0 Cntr1 4 2 0.1 Glucose-1~hosphate:DNA (wlw) 0 1 ~----------~ _0.6 10 5 .. 0.4 0 3 0 0 2 0 1 0 Cntr1 4 2 Sucrose:DNA (wlw) 0 1 0 1 ~-------------~ 0 6 10 5 .. 0 4 c( 0 3 z 0 0 2 0 1 0 0 Cntr1 4 2 Urea:DNA (wlw) 0 1 Figure 8: Effect of lyophilization (FD) on plasmid DNA activity, with various amounts lyoprotectant to DNA (w/w ) Average SEM. n=5 for all treatments, *=FD DNA significantly lower than Control DNA. p<0.05 via Schefe's multiple comparison T-test. 5 hours. The media was then replaced with serum containing media and incubated a further 48 hours before being lysed.

PAGE 59

47 Both forms of luciferase were then assayed using a luminescence spectrophotometer (Monolight 2010). The biofunctionality assay demonstrated a loss of more than 75% of plasmid DNA activity after lyophilization (Figure 8). Furthermore, this loss of activity could be prevented by the use of lyoprotectant carbohydrates (approximately 4:1 carbohydrate to DNA w/w), including glucose, glucose-1-phosphate, glucosamine, lactose and sucrose). These carbohydrates were chosen to study the effect of charge and size on the effectiveness of the lyoprotection. The presence of lyoprotectant did not significantly alter the apparent plasmid DNA concentration either before (data not shown) or after freeze-drying. The protective effect of the glucose-1-phosphate was unexpected if the lyoprotective effect was due to the replacement of the sphere of hydration by the carbohydrate, as it might be assumed that this anionic carbohydrate would be electrostatically repulsed from the major groove by the anionic phosphate backbone. Furthermore, glucosarnine, which would be expected to interact strongly with plasmid DNA, did not exert a statistically significant protective effect at a 4:1 w/w ratio. If the lyoprotectant carbohydrates are protective of plasmid DNA due to

PAGE 60

48 replacement of the water sphere of hydration, then the use of a water destabilizing compound such as urea should decrease DNA activity. However, as seen in Figure 8, urea did support a protective efrect through an undetermined mechanism. Disaccharides demonstrated a greater protective effect on a molar basis than did monosaccharides, suggesting that the size of the protective compound may exert and influence. Potentially, the effect could be due to a non interactive covering of the plasmid DNA that inhibits any conformationally induced denaturation. This hypothesis is supported by the protective effect seen upon ~saturating'' concentrations of all lyoprotectants tested (4000:1 w/w). The extent of interaction between DNA and the lyoprotectant carbohydrates was assessed using thermal analysis to determine onset of melting temperatures. First, behavior in the solid phase was examined using differential scanning calorimetry, a sample DSC scan can be seen in Figure 9. Equal weights of carbohydrate and salmon-sperm DNA were mixed and lyophilized. The lyophilized cake was then assayed to determine the onset of melting. Second, the effect of lyophilization upon melting temperature after rehydration was analyzed using UV spectroscopy. In the dried state, there was a general

PAGE 61

49 trend towards increased onset of melting temperature in the presence of lyoprotectants Figure 10, with a change in the onset of melting suggesting an interaction between the carbohydrates and the DNA. However, glucose-1-phosphate did not show a significant increase in Tm, but did protect against loss of bioactivity, which may suggest that a physical interaction is not necessary for a protective effect. 1 5,---.---~---r----,----.-----.---~--..-----, 5 -~ 5 0 51 4 C 12.68 In -3 08 al C E 4 !i -5 E ...:. v) 0 7 5 -10 1 r, 5,___......_ __._ __ _.__ _._ ___,,___....__ __._ __ ..__ _J -15 1 1 0 8 128 3 145 8 -11 7 5 8 23 3 40 8 58 3 75 8 93 3 unlvers1tv Of Flor10B-MSE TEMP C IHeet1nol u C/) 0 0 Figure 9: Representative thermal analysis scan of salmon sperm DNA. The effect of rehydration of lyophilized plasmid DNA on the DNA melting profile, as measured via absorbance at 260 nm, can be seen in Figure 11. Lyophilized-rehydrated

PAGE 62

50 plasmid DNA showed an earlier onset of melting than plasmid DNA that was kept in solution. This appears to agree with the lower onset of melting seen in the solid state without lyoprotection. It can be seen that the previously lyophilized DNA solutions showed an increased absorption over the 25-80C interval, indicative of the presence of a small amount of single-stranded DNA in the DNA solutions (Lindhal and Nyberg 1972). 60 0 -~ E 20 0 Figure 10: Effect of lyoprotectants on the melting temperature of salmon-sperm DNA. Mean SEM, n=3, = significantly different than unprotected control DNA (p < 0.05) via Fisher's PLSD. To identify a potential mechanistic explanation for the results obtained by the biofunctionality assay, several structural assays were also utilized. Standard agarose gel electrophoresis (0.8% agarose} is capable of separating the conformations of plasmid such as supercoiled, relaxed

PAGE 63

51 circular and linear, by utilizing size fractionation in the agarose gel via an electric field. The DNA is negatively charged because of its phosphate backbone, and will migrate toward the cathode when an electric field is applied. No apparent change in the ratio of supercoiled to relaxed plasmid DNA was observed using this method, Figure 12 and Figure 13, suggesting that the lyophilization process did not cause any gross conformational changes to the plasmid DNA. 0 c.o 0 6 0 5 0 4 0 3 0.2 0 .1 0 0 -l-a~~ ,....Jl t:::::..::~------=-,-=-------=~:!:...._-,-----_J 25 0 3 5 0 4 5. 0 55 0 65 0 7 5 0 8 5.0 Temperature (C) Figure 11: Effect of lyophilization and subsequent rehydration on the melting of plasmid DNA Mean~ SEM, n=3 The standard agarose gel method can not differentiate between undamaged supercoiled and damaged supercoiled plasmid DNA. This damaged supercoiled plasmid DNA could potentially be seen after glycosidic bond cleavage at

PAGE 64

52 ...... w .. -: . .:" ... --~ ... Figure 12: Lyophilized plasmid DNA samples, rehydrated in DI water and run an agarose gel. Samples: marker, HindII digested lambda phage marker; FD, lyophilized DNA without DMED treatment; FD DMED, lyophilized DNA with DMED treatment; pH 3, DNA incubated in pH 3.0, 2.5 rnM citrate buffer for 15 minutes at 50C with no DMED treatment; pH 3 DMED, DNA incubated in pH 3.0, 2.5 rnM citrate buffer for 15 minutes at 50C with DMED treatment before running on gel. abasic sites. Abasic sites are one of the most common DNA lesions and can be produced in at least two ways (McHugh, and Knowland 1995): via spontaneous hydrolysis of the glycosyl bond between deoxyribose and the purines, or at a slower rate between deoxyribose and pyrimidines. These modifications can be accelerated by base modifications such as the alkylation of purines, saturation of C5-C6 bond of the pyrimidines, and fractionation of the heterocyclic ring (Talpaert-Borle, and Liuzzi 1983). In addition, they can be detected by the use of the DMED assay, which cleaves the

PAGE 65

53 phosphate backbone by a beta elimination reaction in the presence of an abasic site (Figure 2 on page 7). < 80 % z C C. ,, 60 % .!! 0 C1) C. 40 % ::, en 0 20 % 0 % 0 1 2 3 Lyophilization Cycles Figure 13: Effect of lyophilization on plasmid form. Mean+ SEM, n=3. p > 0 05 via ANOVA After incubation in the presence of DMED damaged DNA would be expected to show a lower percentage of supercoiled DNA. The percentage of supercoiled DNA did not change after lyophilization and subsequent DMED treatment (Figure 12). This suggests that the lyophilization process did not cause an increase in plasmid DNA abasic sites Analysis by UV/Vis spectroscopy showed a hyperchromic effect for lyophilized plasmid DNA, suggesting potential denaturation or conformational change Any process which increases the interaction of purine and pyrimidine rings, such as a contraction of the plasmid DNA macromolecule or

PAGE 66

54 restriction of internucleotide rotation by the formation of hydrogen bond stabilized helical structures, would result in a hyperchromic effect at 260 nm, as would separation of the two anti-parallel strands (denaturation) (Michelson, 1963). Lyophilization of plasmid DNA resulted in a significant increase in absorbance at 260 nm. This effect can be alleviated by the use of lyoprotectant carbohydrates at a 50 rnM concentration (Figure 14). These results suggested that the lyophilization process causes a conformational change or denaturation in plasmid DNA that can be prevented by the use of carbohydrates as lyoprotectants, since these data agree with the UV melting profile which also suggests a denaturation. 15 0 X E C 10 0 co N 0 .Q 5 Figure 14: Effect of lyophilization on plasmid DNA absorbance at 260 nm. Average+ SEM. n=4, FD DNA significantly higher than lyoprotected DNA p<0.05 via ANOVA

PAGE 67

55 To further identify possible conformational changes circular dichroism analysis was utilized Lyophilized plasmid DNA was compared to "normal" B-form DNA (pH 7.4 TE buffer} and to high-salt, A-form DNA (5 M NaCl} (Bailleal et al 1984; Nishimura et al 1985). As can be seen in Figure 15, the high salt conditions seen during lyophilization cause a conformational change that may be indicative of a change from the B-form to the A-form. 6 ,--.._ ...... 5 FD 0 4 ('I s 3 0 bO V 2 "O '-" w 1 0 250 260 27 0 280 290 3 00 Wavelength (nm) Figure 15 : Wavelength circular dichroism scan of lyophilized plasmid DNA (FD} compared to A-form and B-form plasmid DNA Each scan is the average of three separate baseline subtracted scans The use of lyoprotectants can exert further conformational changes during the lyophilization process Figure 16 The smallest uncharged protectant, glucose, appeared to undergo the least conformational change during the lyophilization process Increasing the size of the

PAGE 68

56 protective molecule, by utilizing lactose as a protectant causes a slight decrease in conformational shift, but not nearly as great a shift as the smaller glucose monomer. When charged moieties were added to the glucose protectant, a further shift in conformation was noted. Both anionic and cationic compounds caused a shift in conformation i3 0 g 2.5 M .., 2 0 J 1 5 ... 8 1 0 05 0 0
PAGE 69

57 solution control state. Furthermore, there is a lack of a clear dose-response relationship between the amount of the carbohydrate and the conformational shift for all compounds except glucose-1-phosphate. It seems that the anionic glucose-1-phosphate requires a 5:1 (w/w) protectant:DNA ratio before the protectant exerts a greater conformational change upon the DNA. This increase in CD measurement is typical of a change in both the helix winding angle and the base pair twist of the plasmid DNA. An increase in the CD measurement signals a shift towards the A-form of the plasmid DNA, with an a decreased winding angle, angle between two adjacent bases, leading to more base pairs per turn of the double helix (Johnson et al. 1981). The kinetics of the conformational change were examined using CD. Essentially, DNA was lyophilized and consequently rehydrated. CD spectra were recorded for multiple time points after rehydration (Figure 17). A first order fit of the data suggests that most of the plasmid returns to the B-form in approximately five hours. At 24 hours after rehydration the measured CD was not statistically different from control non-lyophilized DNA (data not shown), this suggests that the conformation of the plasmid DNA may return to the B-form within 24 hours.

PAGE 70

58 It was hypothesized that this initial conformational change could be responsible for the apparent loss of plasmid DNA activity after lyophilization. Since it was shown that the plasmid returns to the B form in a relatively short period of time plasmid DNA was lyophilized and rehydrated immediately before transfection or rehydrated 24 hours before transfection to determine the A-form effect on plasmid DNA activity (Figure 18) There was no significant difference between O and 24 hours rehydration, suggesting that this potential conformation of the plasmid DNA does not have a direct impact on biological activity. 0 8 0 7 0 6 .., o "'"' 0 5 >< 0) 0.4 C 0 3 0
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59 Another mechanism of damage may be oxidative damage of the plasmid DNA during the lyophilization process. To examine the possibility of this mechanism, plasmid DNA was lyophilized up to six times and then chemically hydrolyzed to the individual bases. The bases were then analyzed by 0 7 ~-------------------~ 0.6 W 0.5 t 0.4 0 3 z C 0.2 0.1 0.0 -1--__J-------.-----.......... L-----,--0 24 Rehydration Time (Hours) Cntrl Figure 18: Effect of rehydration time on plasmid DNA activity. Mean+ SEM. n=6. No significant difference between 0 and 24 hours by T-test. HPLC using parallel UV and ECD detection to quantitate both normal and oxidized bases, specifically guanine and 8-OH guanine. The 8-OH-guanine is a commonly used marker for oxidative damage of DNA. The lyophilized samples showed no significant increase in 8-OH-guanine per mole of guanine and all lyophilized samples were significantly less damaged than the positive control (Figure 20). The type of tube used for lyophilization of plasmid DNA could potentially change the biological activity of

PAGE 72

60 plasmid DNA (Figure 21). If the loss of biological activity was due to the interaction of plasmid DNA and the container used for lyophilization, then the use of ., a, ,.: ., ., "' A "' ., B Figure 19: HPLC analysis of hydrolyzed pRL-CMV plasmid DNA. Panel A is a UV analysis. Peaks by retention time: 2.76, solvent front, 3.79 cytosine, 7.85 guanine, 11.15 thymine, 13.00 unidentified, 17.89 adenine. Panel Bis an ECD analysis. Peaks by retention time 2,67 3.21 solvent front, 3.53 5-OH cytosine, 3.82 cytosine, 4.78 plasticizer, 7.35 guanine, 7.92 8-OH guanine, 16.99 8-OH adenine and adenine. siliconized tubes would be expected to decrease DNA loss. However, there is no significant difference in DNA activity between siliconized and non-siliconized polypropylene tubes, suggesting that plasmid DNA does not interact with the polypropylene tube during the lyophilization process. This is supported by gel electrophoresis, which showed no significant loss of plasmid DNA after cycles of

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61 140 (!) 120 0 E 100 0 0 0 80 .... (!) 60 :::c q 40 00 0 E 20 * 0 0 2 4 6 Cntrl Freeze-drying Cycles Figure 20: Effect of lyophilization of plasmid DNA on oxidative damage. Positive control was Fe + 3 catalyzed oxidation. Mean~ SEM, n=5, except control n=3. = p < 0.05 via Fisher's PLSD versus control. lyophilization. There did appear to be a significant interaction between polyethylene and plasmid DNA during the lyophilization process, suggesting that polyethylene tubes are not an appropriate choice for storage of plasmid DNA. Conclusion In conclusion, lyophilization of plasmid DNA causes a loss of plasmid DNA functionality that can be prevented by the use of carbohydrates as lyoprotectants. The mechanism behind this loss of activity does not appear to be due to a gross structural change, as would be evidenced via agarose gel electrophoresis. Nor does the damage appear to be due to an increase in plasmid DNA abasic sites, as measured by

PAGE 74

62 the DMED assay, increase in oxidative damage as measured via HPLC, or interaction with the enclosure used for E c, 0.7 S 0 6 (.) C 0 5 0 0
PAGE 75

63 protectants were effective against the hyperchromic effect. This suggests that lyoprotectants may decrease a conformationally-induced denaturation of the plasmid DNA caused by the lyophilization process.

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CHAPTER 4 CHARACTERIZATION OF ENDOTOXIN AND CATIONIC LIPOSOME INTERACTION Introduction Gene therapy is one of the fastest growing areas in therapeutics. While rapid progress has occurred in this field, the pharmaceutical community has not adequately addressed safety concerns. In particular, there exists a potential toxicity from plasmid DNA contaminates that could be exacerbated by conventional non-viral gene delivery methods. Since plasmid DNA is typically produced by gram negative bacteria E. coli, endogenous bacterial products can potentially contaminate the final plasmid DNA product (Weber et al. 1995). Typical problematic agents in plasmid DNA preparations include endotoxin (also known as lipopolysaccharide or LPS), a cell wall component that is pyrogenic in man, and DNase, a ubiquitous bacterial enzyme that can degrade the final plasmid product and act as an immunogen. Endotoxin ranks among the most serious limitations in the development of gene products for gene therapy because 64

PAGE 77

65 of its extreme toxicity. E. coli endotoxin consists of a polysaccharide component and a covalently bound lipid component, lipid A. Lipid A is the biologically active component responsible for pathophysiological effects including fever, hypotension, intravascular coagulation and potentially death (Fletcher, and Ramwell 1980; Aida, and Pabst 1990; Rietschel et al. 1993; Xing et al. 1994). Endotoxin is an extremely potent toxin with an LD so of 3 mg/kg and 1 mg/kg in rats and dogs, respectively (Fletcher, and Ramwell 1980; Shibayama et al. 1991). It has been shown that as little as 150 g of endotoxin can be lethal to a horse (Bottoms 1982). Therefore, the Food and Drug Administration (FDA) has established a guidance on human maximal endotoxin dose permissible for parenteral products (F.D.A 1985). This limit is based on endotoxin activity (EU), and can be measured via the LAL (Limulus amebocyte lysate) assay (Levin, and Bang 1964; Levin, and Bang 1968; Iwanaga 1993). Non-intrathecal parenteral drug products are limited to 5 EU kg1 hr1 In the case of an intrathecal parenteral product, the injection limit drops to 0.2 EU kg1 h 1 r These allowable levels are of concern in gene therapy, where the extent of endotoxin contamination in plasmid DNA preparations after treatment to remove endotoxin has been shown to range from 3-15 EU/mg plasmid

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66 DNA, with 1 EU~ 1 ng pure endotoxin (Cotton et al. 1994). Determinations in our laboratory have shown that before specific removal, endotoxin contamination of plasmid DNA can typically approach 5,000 EU/mg, while the literature reports endotoxin concentrations exceeding 15,000 EU/mg DNA (Montbriand, and Malone 1996). It was previously shown that increasing levels of endotoxin contamination decrease transfection efficiency when using DOTAP : DOPE (dioleoyl glycero trimethylammonium propane : dioleoyl glycero phosphoethanolamine) cationic liposomes (Weber et al. 1995) or adenovirus particles (Cotton et al. 1994) to deliver plasmid DNA. However, the mechanism underlying this decreased transfection efficiency has not been clearly defined. It can be predicted that endotoxin interacts with cationic liposomes electrostatically due to the positive charges on the cationic liposomes and the negatively charged phosphorylated glucosamine residues of the lipid A moiety (Rietschel et al. 1993). There also may be other lipophilic methods of interaction with cationic lipids due to the saturated fatty acid chains on lipid A (Rietschel et al. 1993). Upon this interaction of endotoxin, DNA and cationic liposomes, the liposome complex would be expected to be taken into the cell via endocytosis with subsequent

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67 destabilization of the endosome and release of the DNA and endotoxin into the cytoplasm (Xu, and Szoka 1996). It will be of importance for formulation scientists to understand the potential modes of interaction between endotoxin and cationic liposomes. With this knowledge it should be possible to develop newer methods of endotoxin removal from these products. In this report we have focused on endotoxin, similar concerns could be raised with other anionic compounds. Materials and Methods Plasmid Purification A pGL3 plasmid (Promega, Madison, WI) encoding for luciferase driven by the SV-40 promoter was propagated in E. coli JM109 cells (Promega, Madison, WI). The transformed bacteria were cultivated in Lura Bertina (LB) broth containing 100 g/ml ampicillin to select for lactamase encoding plasmid. The pGL3 plasmid was isolated via an alkaline lysis method and purified using an anionic exchange column (Qiagen, Chatsworth, CA). The plasmid was stored in TE buffer (Tris HCl, 10 rnM; EDTA, 1 rnM, pH 7.4). Endotoxin content of the plasmid DNA before endotoxin removal was approximately 4,000 EU/mg, as determined by the LAL assay method {QCL-1000, BioWhittaker, Walkersville,

PAGE 80

68 MD). Endotoxin was removed by the Triton X-114 method of Manthorpe, et al (Manthorpe et al. 1993). Liposome Preparation Lipids were obtained from Avanti Polar Lipids (Alabaster, AL). Cationic DOTAP : DOPE, NBD labeled DOTAP : DOPE, and zwitterionic lecithin liposomes were prepared using the hand-shaking method (New 1990). DOTAP: DOPE (10 mg DOTAP, 10 mg DOPE), NBD labeled DOTAP: DOPE (10 mg DOTAP, 10 mg DOPE, 0.1 mg 1-palmitoyl-2-[12-[ (7-nitro-21,3-benzodiamino]dodecanoyl]-sn-glycero-3-phosphate), or lecithin (10 mg) was added to 10 ml of chloroform and introduced into a 250 ml round bottom flask. Chloroform was evaporated using a rotary evaporator at 60C. The lipid film was hydrated with 10 ml of distilled water and shaken for 30 minutes at 60C. Lipids were sized by extrusion 6 times through 200, 600, or 800 nm polycarbonate filters (Poretics Corp, Livermore, CA). Sizes were confirmed with a laser light scattering particle sizer using volume weighted distribution (Nicomp 380ZLS, Santa Barbara, CA). Transfection Efficiency Assay Phenol purified, unlabeled endotoxin (0-50,000 EU/ml), E. coli serotype O55:B5 (500 EU/g, Sigma Chemical Company,

PAGE 81

69 St. Louis, MO) was added to a pGL3 luciferase plasmid: cationic lipid mixture (DOTAP DOPE 1:1 w/w) with a ratio of 1 g plasmid to 2 g DOTAP. The mixtures were incubated for 15 minutes to allow interaction of the anionic plasmid to the cationic liposomes, followed by transfection into COS-1 cells (ATCC, Rockville, MD) that had been plated 24 hours earlier at 3xl0 4 cells/ml in 24 well tissue-culture dishes. All transfections were carried out using serum free DMEM (Dulbecco's Modified Eagle's Medium) supplemented with penicillin G (100 units/ml) and streptomycin (100 g/ml). They were subsequently incubated for 5 hours in a humidified, 5% CO 2 incubator at 37C. The media was aspirated and the cells washed with PBS (phosphate buffered saline) before being replaced by 10% FBS (fetal bovine serum) containing DMEM and antibiotics. The cells were incubated for an additional 19 hours, washed with 1 ml PBS, and then lysed with lysis buffer (200 L, 0.1 M potassium phosphate, 1% Triton X-100, 1 mM DTT, 2 mM EDTA, pH 7.8). Luciferase activity was determined using a luminescence spectrophotometer (Monolight 2010, Analytical Luminescence Laboratory, San Diego, CA). Lysate (20 L) and the assay buffer (100 L, 30 mM tricine, 3 mM ATP, 15 mM MgSO 4 10 mM DTT, pH 7.8) were added to the sample cuvette.

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70 D-luciferin (100 L, 1 rnM, pH 6.5) was injected to initiate the reaction. The light output was measured for 10 seconds and the results integrated to yield the luciferase activity. All luciferase measurements were standardized against protein concentration as measured by the bicinchoninic acid protein assay (BCA Protein Assay, Pierce, Rockford, IL). Anisotropy Assay A constant amount (1000 EU) of flourescein isothiocyanate (FITC) labeled endotoxin, E. coli serotype O55:B5 (100 EU/g, Sigma Chemical Company, St. Louis, MO), and varying amounts of unlabeled cationic DOTAP: DOPE liposomes (600 nm, 0-16 g), zwitterionic lecithin liposomes (200 and 800 nm, 0-16 g,), PAMAM (polyamidoamine) dendrimer (generation 4, 0-16 g, in TE buffer, pH 7.4, Aldrich Chemical, Milwaukee, WI), or plasmid DNA (pGL3 in DI water), with a final volume of 1 ml, were allowed to 0 equilibrate together for 15 minutes at 25 C. Fluorescent 0 anisotropic measurements were conducted at 25 C, 487 nm and 525 nm excitation and emission wavelengths, respectively using a luminescence spectrophotometer (LS50B, Perkin Elmer, Oak Brook, IL).

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71 Endotoxin Dephosphorylation Unlabeled endotoxin (30 EU) was incubated for various time from Oto 120 minutes with 1 mU alkaline phosphatase (Promega, Madison, WI) at 37C following the methods of Poelstra, et al (Poelstra et al. 1997a; Poelstra et al. 1997b) with minor variation in the assay buffer: 0.5 M Tris, 10 mM MgC1 2 1 mM ZnC1 2 pH 7.8. Endotoxin Assay Endotoxin activity was quantitatively measured using a commercially available chromogenic LAL test method (QCL1000, BioWhittaker, Walkersville, MD) according to the manufacture's instructions. In brief, endotoxin (0.6 EU) was incubated for 15 minutes with an increasing concentration of DOTAP: DOPE cationic liposomes (0-100 ng) and brought to a final volume of 50 L with endotoxin free water. The LAL reagent (50 L) was added to the reaction vessel and the mixture was further incubated for 10 minutes at 37C. Chromogenic substrate solution (100 L) was added and the sample incubated for 6 minutes before the addition of stop reagent (100 L, 25% glacial acetic acid in water). Absorbance of the p-nitroanaline product was read at 405 nm using a UV/Vis spectrophotometer (Lambda 3, Perkin Elmer, Oak Brook, IL)

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72 Cell Viability Assay The MTT (dimethylthiazol diphenyltetrazolium bromide) assay (Freshney 1994) was used to assess the effect of the endotoxin and liposome delivery systems on cell viability. COS-1 cells were plated in a 24 well tissue-culture plate at a density of 3xl0 4 cells/well in 1 ml DMEM media containing 10% FBS and incubated for 12 hours in a 37C, humidified, 5% CO 2 incubator. The serum containing media was aspirated, washed with PBS, and replaced with serum free media. Varying amounts of lipid (0-170 g), endotoxin (0-50,000 EU), and lipid: endotoxin combinations (5 g: 050,000 EU) were added to each well and incubated for 5 hours. The media was then changed back to 10% FBS DMEM and the incubation continued until 24 hours following the addition of test formulations. The cells were then fed with 1 ml of fresh media and MTT (250 L, 5 mg/ml) and were incubated for 5 hours and then the media was removed. Dimethylsulfoxide (DMSO, 1 ml) and glycine buffer (250 L, 0.1 M glycine, 0.1 M NaCl, pH 10.5) were added and the absorbance was immediately read at 570 nm. Untreated cells and buffer alone were used as positive and negative controls, respectively.

PAGE 85

73 Statistical Analysis Statistical analysis between the various treatments was conducted using analysis of variance and Fisher's protected least significant difference (PLSD) post-hoc tests where appropriate (StatView v4.5, Abacus Concepts, Berkley, CA), with p<0.05 considered statistically significant. Results The effects of endotoxin contamination upon transfection efficiency were determined by adding increased levels of endotoxin to pGL3 luciferase plasmid: cationic lipid mixture (DOTAP: DOPE 1:1 w/w) with a ratio of 1 g plasmid to 2 g DOTAP before transfection into COS-1 cells (Figure 22). A 1:2 ratio of plasmid to cationic lipid results in a neutral to slightly negative net charge for the overall complex. In the presence of DOTAP: DOPE cationic liposomes (2 g/ml), low levels of endotoxin (50 EU/ml) appeared to increase the variability of luciferase reporter activity, a marker of the plasmid DNA functionality. Higher concentrations of endotoxin resulted in a corresponding decrease in the luciferase reporter activity. Activity decreased more than 90% at 5000 EU/ml

PAGE 86

74 endotoxin, however the cells appeared viable under direct observation. 50 -v 40 0 X C 30 2 e a. o 20 E -::::, _J 10 0::: ....... Liposome No Liposome 0 50 500 5,000 Endotoxin (EU/ml) 50,000 Figure 22: Enzyme activity corrected for total cellular protein after transfection of luciferase plasmid (lg) in the presence of endotoxin, with and without DOTAP: DOPE cationic liposomes. with lipid (2 ug/ml), without lipid. RLUSEM, n=4, p<0.05 via one way ANOVA for effect of endotoxin in the presence of cationic lipid. The extent of interaction between endotoxin and cationic liposomes (DOTAP: DOPE) was determined using fluorescence anisotropy. The potential contribution of electrostatic versus lipophilic interactions was investigated by varying the ionic strength of the incubation solution (NaCl, 0-2 M). There was a correlative increase in the anisotropic measurement when endotoxin was incubated with additional cationic lipid, suggesting formation of a complex (Figure 23). Maximum binding occurred at a 1:2 (w/w) ratio of total lipid to endotoxin.

PAGE 87

75 The interaction decreased with increasing ionic strength, but the size of the complex increased significantly, as monitored by change in anisotropy, even under high ionic strengths (2 M NaCl), suggesting the presence of electrostatic in addition to other interactions. Free FITC was used as a control to ensure the interaction detected was due to endotoxin and not a consequence of the FITC label (Figure 23 inset). Furthermore, the spectral properties of the free FITC and the conjugated FITC appear similar as determined by excitation and emission maxima. The potential for competition between plasmid DNA and endotoxin for DOTAP : DOPE cationic liposome was compared using a similar experimental paradigm. DOTAP: DOPE (5 g) and FITC conjugated endotoxin (1000 EU) were held constant at the maximal binding ratio seen in Figure 23. As plasmid DNA levels were increased, there was a decrease in anisotropy readings, suggesting increased competition between endotoxin and plasmid DNA for interaction with DOTAP: DOPE. This results in the displacement of endotoxin (Figure 24). To further characterize the mechanism of endotoxin and cationic liposome interaction, additional fluorescence anisotropy studies were carried out. Two sizes of PAMAM dendrimers, a cationic cascade polymer, were studied. The

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0.12 0.10 C 0.08 >, a. e 0.06 ..... 0 !!? C <( 0.04 0.02 0.00 0 2 4 76 0 12 .-------~ :S 0 10 i>: 0 08 _g 0 06 -~ 0 04 0 02 FreeFITC 0 00 L.....t=;::l=~::::::::::::::J 6 8 10 12 DOTAP:DOPE (g) 14 16 ...,. DI Water 0.5 M NaCl ...._ 1 M NaCl -M-2MNaCI Figure 23: Effect of increasing DOTAP: DOPE liposomes concentration with FITC-endotoxin held constant (1000 EU) on anisotropy (r) DI Water, O. 5 M NaCl, A 1. 0 M NaCl, X 2.0 M NaC l Inset: Free FITC (22 ng) 1.0 M NaCl. Change in r+SEM, n=3, p<0.05 using two way ANOVA for increase in r with increasing lipid under all conditions. dendrimers would be expected to interact with FITC labeled endotoxin though electrostatic forces since the dendrimers only exposed functional groups are amines. Also, two sizes of zwitterionic lecithin liposomes (0.2 and 0.8 m) were examined to determine the importance of the other interactions between the lipids. The greatest change in anisotropy occurred with the cationic PAMAM dendrimers (Figure 25). Smaller, though statistically significant, changes in anisotropy were seen at the same concentration of lecithin, suggesting that the prevalent mechanism of

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77 0.10 -c0.08 ._,, >, a. 0 'E 0.06 (/) <( 0 04 0.02 0 2 4 6 8 pDNA (g) 10 12 14 16 Figure 24: Effect of increasing luciferase plasmid concentration on anisotropy (r) with constant FITCendotoxin (1000 EU) and DOTAP : DOPE liposomes (5 g). Change in rSEM, n=3, p<0.05 using two way ANOVA for decrease in r with increasing plasmid. interaction is electrostatic rather than lipophilic. In the case of both sizes of dendrimers and liposomes, the larger sized particles led to a greater change in anisotropy as expected, due to the additive nature of anisotropy. Further evidence for interaction between endotoxin and cationic liposomes was demonstrated using a gel retardation assay (Figure 26), which confirmed the

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78 FITC-endotoxin cationic lipid complex by retarding the progress of the endotoxin band in an agarose gel. 0.1 0.1 0.1 S 0.1 >, a. e 0 0.0 .!!l C: <( 0.0 0 0 0.0 gadded .,... Dendrimer G4 Dendrimer G2 -6-Liposome 0.8m -Liposome 0 2m Figure 25: Effect of increasing lipid or dendrimer concentration with FITC-endotoxin held constant (1000 EU) on anisotropy (r). Dendrimer (Generation 2), Dendrimer (Generation 4), X lecithin liposomes (0.2 m), A lecithin liposomes (0 .8 ) Change in rSEM, n=3, p<0.05 using two way 'ANOVA for increase in r with increasing concentration under all conditions. Well LPS 0 3 6 8 10 12 14 16 DOTAP:DOPE (g) Figure 26: Effect of increasing DOTAP: DOPE liposome concentration with FITC-Endotoxin held constant (10 g).

PAGE 91

79 The effect of dephosphorylation of endotoxin on cationic liposome interaction was also examined (Figure 27). Unlabeled endotoxin (30 EU) was incubated for varying time with calf intestinal alkaline phosphatase. The dephosphorylated endotoxin was allowed to interact with fluorescent NBD labeled DOTAP: DOPE cationic lipid (10 g, 600 nm). Increased incubation time with the alkaline phosphatase led to a trend towards decreased anisotropy (p=0.2 via ANOVA). The anisotropy signal seen after 120 minutes of endotoxin incubation was statistically equivalent to labeled lipid alone. The small change in anisotropy was due to the label attached to the liposome and to the high ionic strength of the reaction buffer, resulting in a small size change upon interaction. The chromogenic LAL assay was used to determine the effect of interaction of endotoxin and cationic lipid upon the toxicity of the lipid A moiety. All determinations were compared to a standard curve of known endotoxin concentrations. No endotoxin activity was evident with the cationic lipid alone. The LAL assay showed no significant change in the endotoxin activity when free endotoxin was compared with cationic lipid-endotoxin complex (Figure 28).

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... 0.015 >Q. 0 ... 0 II) C 0.010 <( 0.005 0 20 40 80 60 Time (min) 80 100 120 Figure 27: Effect of increasing incubation time of alkaline phosphatase and unlabeled endotoxin (33 EU) on anisotropy (r) with constant and NBD labeled DOTAP: DOPE liposome (10 g). Change in r SEM, n=4, p>0.05 via one way ANOVA for decrease in anisotropy over time. Literature reports suggest that cationic lipids are toxic in vitro and in vivo (Scheule et al. 1997); however the toxicity of the cationic lipid endotoxin complex is not known. The MTT assay was used to determine the effect of the complex on COS-1 cell viability. Using this method, a dose-response curve was first determined for the DOTAP DOPE cationic lipid. Dose-response curves were then examined using endotoxin alone and endotoxin incubated with

PAGE 93

81 relatively nontoxic concentrations of lipid (<5 g/ml). While the DOTAP: DOPE ......... 0 8 E ::> w >0 6 ..... > +J u <( C x 0.4 0 ..... 0 -0 C w 0 2 0 .1 1 10 10 0 DOTAP : DOPE (ng) Figure 28: Effect of increasing lipid concentration with endotoxin held constant (0 6 EU) on endotoxin activity, EU/ml+ SEM, n=3, p>0.05 via one way ANOVA for effect of cationic lipid on endotoxin activity. cationic lipid was toxic, no significant loss of COS-1 cell viability was detected at endotoxin concentrations up to 5000 EU/ml in the presence of lipid and no change in viability was seen with free endotoxin (Figure 29). Discussion The interactions of endotoxin with non-viral gene delivery systems and the resultant decreases in

PAGE 94

82 transfection efficiency are of potential concern in the success of gene delivery. If a fundamental understanding of the factors that influence endotoxin-delivery system interaction is gained, vectors might be developed that limit the impact of endotoxin in these formulations. 120 A B 100 m 80 > '60 0 40 20 0 1 10 100 5 50 500 5000 DOT AP: DOPE (ng/ml) Endotoxin ( g/ml) Figure 29: Effect of increasing cationic lipid (panel A) and endotoxin (panel B) on COS-1 cell survival via MTT assay. mean~ SEM, n=5, p < 0.05 via ANOVA for DOTAP: DOPE, p > 0.05 for endotoxin via ANOVA. Fluorescent anisotropy results demonstrate endotoxin interacts with DOTAP : DOPE cationic liposomes. Fluorescent anisotropic measurements are well suited for observing interactions between molecules since the reporter fluorescent probe is sensitive to its environmental conditions. Fluorophores have a defined orientation and preferentially absorb light that is vectored in that

PAGE 95

83 orientation. By excitation using polarized light, it is possible to selectively excite individual molecules. As molecules undergo faster rotational diffusion in solution, the time of fluorescence emission and anisotropy decreases. As the molecular volume increases with interaction, there is a correlative decrease in rotational diffusion and thus an increase in the value of the anisotropic measurement. This anisotropic methodology has been applied to observing oligonucleotide hybridization in solution (Murakami et al. 1991) and to examine oligonucleotide-dendrimer interaction (Poxon et al. 1996). The structure and physical properties of endotoxin contribute to the interaction with cationic DOTAP: DOPE liposomes. For instance the fatty acid chains present on the lipid A moiety contributes to the amphipathicity of endotoxin. Endotoxin's amphipathic nature coupled with its low solubility causes aqueous preparations of endotoxin to exist in mainly an aggregated state (Takayama et al. 1995). The prevalence of the aggregated state results in the clustering of the negative charge. This cluster of charge should help to contribute to the electrostatic interaction between the cationic liposomes and the anionic endotoxin. The LAL assay provided information on the effects of the endotoxin-cationic lipid interaction on potential

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84 biological activity. The interaction of the endotoxin and cationic DOTAP liposomes did not alter the activity of the endotoxin as measured by the LAL assay. This implies either that the interaction does not hinder the toxic lipid A moiety or that the complex is not stable. The MTT assay demonstrated the effects of endotoxin cationic lipid complex on COS-1 cell viability. Increasing levels of endotoxin-cationic lipid complex and free endotoxin had no significant effect upon COS-1 cell viability. This lack of selective toxicity in an immortalized cell line has previously been reported, including HeLa, Vero, 3T3, K562, WI-38, SVl, TX-4, CHO, P3U3, and R-393 cells (Epstein et al. 1990, ; Cotton et al. 1994). Immortalized cell lines have shown no change in growth as determined by doubling time, plating density and confluent density measurements. Rather, the toxic response occurs in primary cell cultures cells (Epstein et al. 1990; Cotton et al. 1994). In any case, the effect of endotoxin in a more complex in vivo system is extremely toxic and should not be dismissed. The mechanism of endotoxin-cationic lipid interaction is thought to be mainly electrostatic, as evidenced by the trend to decreased interaction after endotoxin dephosphorylation and ionic strength experiments. However,

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85 other forces may be involved due to the inability of high screening-ion concentrations to completely inhibit the interaction phenomena and the ability of zwitterionic liposomes to interact with endotoxin It should be noted that there were a different number of particles with a different exposed surface area at similar weights of lecithin liposomes and cationic dendrimers This fact could change the rate of endotoxin interaction with each compound. However, the greater change in anisotropy observed with the cationic dendrimers, which should undergo solely electrostatic interaction is most likely indicative of a greater electrostatic interaction Structural flexibility of the vesicular liposome versus the more static solid dendrimer may also have an impact on the interactions. When endotoxin undergoes an electrostatic interaction with cationic liposomes, this interaction can result in lower transfection efficiencies, as measured by enzymatic expression Furthermore, this effect may not solely be due to a difference in the delivery of the complex as supported by these studies Previous studies with transiently transfected CHO cells show that exposure to endotoxin after transfection results in a stimulation of the transfected product at 10 ng/ml endotoxin but decreases production at higher levels

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86 (Epstein et al. 1990). A trend towards a bimodal response can be seen in Figure 22, but was not statistically significant in this study, resulting only in increased variability. These results suggest that the effect of endotoxin on established cell lines is not through a cytotoxic mechanism but rather a difference in the delivery of the plasmid endotoxin-cationic lipid complex. The inclusion of endotoxin into the cationic liposome complex may alter the morphological form and overall net charge of the complex, thereby altering the delivery and subsequent transgene expression. The displacement of DNA from cationic liposomes by endotoxin is most likely responsible for the decreased transfection efficiency in established cell lines, since lowered delivered plasmid levels would result in decreased expression. From the present study it is clear that several factors besides the plasmid and delivery vector will influence the activity of non-viral gene delivery systems. Endotoxin contamination can potentially impact transfection efficiency via competition with plasmid DNA for cationic liposome binding, but this would not be expected with typical GLP or GMP preparations used in clinical studies where the endotoxin levels range from 10 1000 EU/mg DNA.

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87 At these reduced endotoxin levels, the competition between endotoxin and DNA for the cationic liposome is at best marginal. An effect can be seen, with increased transfection variability, at 50 EU/ml. This is a level of endotoxin contamination that can occur with small scale plasmid preparations used for in vitro cell transfections, and potentially affects the results seen in many in vitro studies.

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CHAPTER 5 FOAM FRACTIONATION AS A METHOD TO SEPARATE ENDOTOXIN FROM RECOMBINANT BIOTECHNOLOGY PRODUCT Introduction Biotechnology based therapy, using products isolated from E. coli, need to be concerned of a potential for toxicity from the contaminant endotoxin (Weber et al. 1995). Endotoxin, also known as lipopolysaccharide or LPS, is a gram negative bacterial cell wall component commonly co-isolated with plasmid DNA and recombinant proteins. It consists of a polysaccharide component and a covalently bound lipid component, lipid A. Lipid A is biologically active and can cause a number of pathophysiological effects including fever, hypotension, intravascular coagulation and death (Fletcher, and Ramwell 1980; Aida, and Pabst 1990; Rietschel et al. 1993; Xing et al. 1994). The removal of plasmid DNA endotoxin contamination can be difficult on several accounts (Cotton et al. 1994). First, the negative charges associated with lipid A will cause endotoxin to mimic DNA on anion exchange resins. Second, the large size of endotoxin molecule aggregates 88

PAGE 101

89 will result in a similar retention behavior for both endotoxin and DNA on a size exclusion column. Third, the comparable densities of endotoxin and DNA will result in contamination of plasmid DNA bands in cesium chloride centrifugation. Finally, endotoxin precipitates from ethanol and isopropanol solutions along with plasmid DNA. Similar problems can occur with recombinant proteins, which can also have a negative charge and size that can result in co-purification of endotoxin. The most reliable way of degrading and detoxifying , 0 0 endotox1n is by dry-heat, 180 C for 3 hours or 250 C for 0.5 hours (Tsuji, and Lewis 1978). However, this method is unacceptable for plasmid DNA and proteins, which could be irreversibly denatured (Stroop, and Schaefer 1989). Another standard method, potassium permanganate oxidation, would also unacceptably oxidize both types of product. Several other methods have been proposed for the removal of endotoxin from DNA. Phase separation, with Triton X-114, has been used to reduce endotoxin levels for both protein (Aida, and Pabst 1990) and plasmid (Manthorpe et al. 1993; Cotton et al. 1994). However, the Triton X-114 method is difficult, especially when purifying large volumes, because of the difficulty of separating the endotoxin containing Triton gel phase from the plasmid and protein containing

PAGE 102

90 aqueous phase. A triple-helix oligonucleotide affinity chromatography method has been employed to reduce endotoxin contamination for plasmid DNA, but would not be effective for protein (Wils et al. 1997). As well, gel and resin based endotoxin affinity chromatography systems have been utilized to remove endotoxin from plasmid DNA and protein preparations (Wicks et al. 1995; Montbriand, and Malone 1996; Tan et al. 1997). The main disadvantage of these methods is the loss of large amounts of product during the endotoxin removal. Ultrafiltration methods commonly used for parenterals can potentially remove the product of interest as well as the contaminating endotoxin because of size constraints. These facts suggest that new, efficient methods of purification be developed. In this report we investigate the use of foam fractionation as a potential method to separate endotoxin from biotechnology products in solution. Foam fractionation separates the components of a solution if they have differing surface activities (Lima, and Varley 1996). Stable foam is produced when gas is sparged through a liquid containing a surface-active compound, producing bubbles. Surface-active compounds are enriched around the bubbles, due to the high surface area. These bubbles rise through the liquid forming foam at the surface. This foam,

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91 when removed and allowed to collapse, is made up mainly of surfactant enriched liquid (Lima, and Varley 1996). This technique has been used for the purification of many proteins including bovine serum albumin, lysozyme, ~-casein (Lima, and Varley 1996), and bacitracin (Oka et al. 1991). For non or weakly surface-active compounds other surfactants have been added to act as foaming agents (Grieves, and Aronica 1966). These pro-foaming agents may be needed for the removal of extremely dilute, but still unacceptably high, levels of endotoxin from plasmid DNA solutions, or to foam recombinant proteins that are weak surfactants. Methods Plasmid Purification A pGL3 plasmid (Promega, Madison, WI) encoding luciferase and ~-lactamase was grown in E. coli JM109 cells (Promega, Madison, WI). The transformed bacteria were cultivated in Lura Bertina (LB) broth containing 100 g/ml ampicillin. The pGL3 plasmid was initially isolated using an alkaline lysis method, subsequently purified on an anionic exchange column (Qiagen, Chatsworth, CA) and stored in TE buffer (Tris HCl, 10 mM; EDTA, 1 mM; pH 7.4). The

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92 resultant DNA contained 2.5 EU/g DNA as determined by the Limulus Amebocyte Lysate assay (QCL 1000, BioWhittaker, Walkersville, MD). Foam Fractionation The foam fractionation experiments were carried out using a cylindrical glass column 50 cm tall and 2 cm inside diameter. Nitrogen gas saturated with water was bubbled through 20 ml of various test solutions at 10, 500 and 5000 ml/min. Test solutions contained FITC conjugated endotoxin (100 g/ml, E. coli serotype O55:B5; Sigma Chemical Company, St. Louis, MO) and were stored at 3 pHs (40 mM citrate, pH 3; 100 mM glycine, pH 10; or DI water pH 6.5). Samples were periodically taken through a port at the bottom of the column. All experiments were continued for 60 90 minutes. The FITC signal from each time point was quantified using a fluorescence plate reader (Fmax, Molecular Devices, Sunnyvale, CA) at an excitation wavelength of 487 nm and an emission wavelength of 525 nm, with a standard curve determined for each experiment. Plasmid and FITC-Endotoxin Gel Analysis Plasmid DNA (25 g/ml) or FITC-endotoxin (100 g/ml) was treated in the aforementioned apparatus for 90 minutes.

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93 An aliquot was then electrophoresed on a 0.8% agarose gel at 50 volts for 2 hours in half strength TBE buffer (50 mM Tris, 50 mM borate, 1 rnM EDTA, pH 7.8). The gel was stained with ethidium bromide for DNA analysis and left unstained for FITC-endotoxin analysis. It was photographed under UV illumination with a DC40 digital camera (Kodak, Rochester, NY), Surface Activity Determination Surface activity was determined at 25C using a DuNoliy Interfacial Tensiometer, model 70545 (CSC Scientific, Fairfax, VA) following the manufacturer's instructions. Particle Size Analysis Particle size analysis was carried out using a laser light scattering particle sizer (Nicomp 380 ZLS, Particle Sizing Systems, Santa Barbara, CA). A volume-weight, vesicle, nongaussian distribution was used in all analysis. Endotoxin Assay The Limulus amebocyte Lysate, LAL, Assay (QCL1000, BioWhittaker, Walkersville, MD) was used to quantify all unlabeled endotoxin following the manufacturer's instructions.

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94 Protein Analysis Bovine serum albumin (BSA, Sigma Chemical Company, St. Louis, MO) was used as a representative protein. Concentrations were determined by comparison of absorbance at 280 nm to a standard curve using a spectrophotometer (Lambda 3, Perkin Elmer, Oak Brook, IL). Statistical Analysis Statistical analysis between the various treatments was conducted using analysis of variance and Fisher's protected least significant difference (PLSD) post-hoc tests where appropriate (StatView v4.5, Abacus Concepts, Berkley, CA), with p<0.05 considered statistically significant. Results and Discussion While the system could effectively separate a test solution of bovine serum albumin and FITC conjugated endotoxin in solution (Figure 30), the foam fractionation method was not effective in separating FITC-endotoxin from plasmid DNA solution. In an attempt to remove endotoxin from plasmid DNA solution by foaming, a pro-foaming agent was examined: Triton X-100 (0.03 and 3 mM). There was no foaming effect seen using either labeled or unlabeled endotoxin without the presence of pro-foaming agents (pH 3,

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95 6.5 or 10 and sparging rates up to 5 L/minute). The addition of Triton X-100 caused foaming both below (0.3 rnM) and just above (3 rnM) its critical micelle concentration (CMC 2 rnM). The addition of BSA (1 g/ml) also caused foaming, with flow rates as low as 10 ml/minute. However, no decrease in the FITC-endotoxin concentration was seen (Figure 30). Unlabeled endotoxin could not be measured after the addition of Triton X-100 as a pro-foaming agent, due to the inhibition of the Limulus Amebocyte Lysate (LAL) assay that is used to determine endotoxin activity. This was most likely due to change in LAL enzymatic activity caused by denaturation after the addition of surface-active pro-foaming agents. In a further attempt to separate plasmid DNA from endotoxin, the ionic strength of the solution was increased by the addition of sodium chloride to a final concentration of 1 M. This resulted in a dramatically smaller bubble size (visual analysis), increasing the available surface area for endotoxin separation because of an increase in the surface tension (Wong, and Parasrampuria 1997). The increase in ionic strength did not have any effect in foaming FITC-endotoxin.

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100 90 80 ::; E 70 -Cl C 60 x 0 0 50 --0 C w I 40 (.) Iu::: 30 20 10 0 96 :I __ _._ __ r ~ ____ ... _____ ! 900 850 800 750 CD (/) 700 .: 650 'e. 3 600 550 500 .c: -<--....----r-----,-------.----,,----r-----.--~450 0 10 20 30 40 50 60 Minutes Fractionated LPS / Triton (3 mM) _._ LPS / Triton (0 03 mM) -A-LPS / BSA (1 mglmL) BSA Figure 30: Effect of foam fractionation on FITC labeled endotoxin levels (initial 100 g/ml n=3) and BSA (initial 910 g/ml, n=3 ) in a plasmid DNA solution, (25 g/ml ) mean + SEM. p>0.05 using one way ANOVA for differences in endotoxin concentration over time. p<0.05 for differences in BSA concentration over time. To ensure that extended foam fractionation does not break down the FITC-endotoxin, samples were fractionated for up to 90 minutes. Then these samples were electrophoresed on a 0.5% agarose gel. No breakdown products were detected for FITC-endotoxin upon electrophoresis. As well, extended foam fractionation had no effect on endotoxin activity, as measured via the LAL assay (Figure 31).

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97 Plasmid DNA (25 g/ml) alone did not foam when sparged with notrogen at flow rates up to 5 L/min nitrogen. In addition no obvious structural breakdown or change in concentration could be detected for plasmid DNA alone, or with Triton X-100 pro-foaming agent, after foam fractionation (Figure 32) These data agrees with prior observations that highly anionic DNA does not foam (Lalchev et al. 1982). 25000 E 20000 ... ... ::::::, & & .... w 15000 C >< 0 10000 .., 0 "C C 5000 w 0 I I I I I 0 10 20 30 40 50 60 Minutes Frationated Figure 31 : Effect of foam fractionation on endotoxin activity. Mean SEM p > 0 05 via ANOVA. The inability to separate FITC-endotoxin from plasmid DNA solution, and consequently the ability to separate endotoxin and surface-active protein, is most likely due to the labeled endotoxin's low surface activity. This can be

PAGE 110

98 Minutes Fractionated Figure 32: Effect of foam fractionation on physical stability of plasmid DNA, molecular weight marker A phage Hind III digest in lane 1; lanes 2 8, fractionation time points 1 g pDNA per lane, in a 0.8% agarose gel stained with ethidium bromide. seen in Figure 33, where FITC-endotoxin causes no significant change in surface tension as compared to distilled water. The FITC label appears to decrease the surface activity as compared to the unlabeled endotoxin. However, as stated previously, unlabeled endotoxin could not be made to foam at flow rates up to 5 L/min nitrogen. As well, the effect of pro-foaming agents on unlabeled endotoxin was unable to be determined because of assay inhibition. Lack of surface activity of FITC-endotoxin and subsequent lack of separation may also be due to the endotoxin molecules tendency to aggregate when in aqueous solution (Takayama et al. 1995). In the aggregated state the endotoxin molecules would be oriented in a micelle so

PAGE 111

80 E ~75 u, Q) C "O --. 70 u, C Q) ..... ~65 ::J (/) 60 99 BSA FITC-LPS LPS Water Figure 33: Surface tension (dynes/cm) of water, endotoxin, FITC-endotoxin, and BSA (all 1 mg/ml), n=4, mean+ SEM, = p<0.05 using Fisher's PLSD versus water. that their lipophilic moieties would be hidden, reducing the apparent lipophilic nature of the aggregate but not the free endotoxin. The tendency for aggregation may have effected the surface activity measurements in Figure 33. It has proven possible to disaggregate endotoxin by altering pH. As FITC-endotoxin solution is acidified, aggregate particle diameter decreases (Figure 34). This disaggregation occurs commensurately with a decrease in surface tension (Figure 35). At acidic pH, protonation of endotoxin may increase the solubility of the monomeric form of FITC-endotoxin. This would result in a shift away from the aggregate form and towards the more highly surface

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100 active monomeric form resulting in a significant but minor decrease in surface tension However, changes in pH did 35 E 30 C -Q) t::! 25 en Q) u t 20 cu a.. Q) ... cu 0) 15 Q) ...... 0) ;f C 10 x 0 ... 0 u C 5 w I (.) ILL 0 1 10 100 1000 HC I ( M) Figure 34 : Volume weighted particle size of FITC-endotoxin (1 mg/ml) at increasing HCl concentration Data are shown as meanSEM., p<0.05 via ANOVA, *= p<0 05 using Fisher's PLSD verus 1 M HCl not result in the foaming of FITC-endotoxin While this result may be discouraging for the separation of endotoxin from plasmid DNA, it is encouraging for the separation of

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101 endotoxin from surface-active recombinant proteins, since pH extremes are sometimes needed to successfully foam protein from solution (Lockwood et al 1997) 80 ---------------------, 79 ......... E (/) Q) C: 78 >. 0 ---C: Q en C: Q) I77 Q) (.) ::::, (J) 76 75 ------r-----.---....----r------r----r---0 200 400 600 800 1000 HCI ( M) Figure 35: Surface tension (dynes/cm) of FITC-endotoxin (1 mg/ml) at various HCl concentrations n=4 mean ~SEM, = p<0.05 using Fisher s PLSD versus O m HCl. A number of changes were made to this system to ensuring that endotoxin does not foam in aqueous solution Variables that normally are altered to increase the foaming

PAGE 114

102 of recombinant proteins were examined to determine their effect on FITC-endotoxin foaming. These included the flow rate of the sparged gas, ionic strength, pH, and the concentration of pro-foaming agent. FITC-endotoxin remained in the bulk solution under all conditions. This is reassuring for the use of foam fractionation as a method to separate surface-active proteins from endotoxin. These studies suggest that foam fractionation will be ineffective in the purification of plasmid DNA solutions, which are not surface-active and remain in the bulk solution with endotoxin. These data support the use of foam fractionation as an effective method for separating surface-active recombinant proteins, which will concentrate in the foam, away from bulk solution endotoxin.

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CHAPTER 6 CONCLUSIONS As stated in the introduction, the overall goal of this project was to determine the impact of environmental factors on plasmid DNA stability. Using structural and biofunctional assays, the stability and mechanisms of plasmid DNA instability were analyzed. This information was then assessed with respect to the pharmaceutical development of plasmid DNA. Four specific hypotheses were investigated by this project. The first hypothesis was that biofunctional assays are more sensitive than structural assays, since damage at one base in the encoding region would not be easily detected by conventional structural methods, but could result in a functionally inactive product. This hypothesis was not shown to be true. For some types of damage, specific structural methods were more sensitive than biofunctional assays. One example of this difference can be seen when plasmid DNA is stored at 37C for three weeks. Functionally, no significant difference was seen between control plasmid DNA and the treated plasmid. However, 103

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104 analysis by the DMED assay showed that there was an increase in abasic sites following prolonged storage at body temperature. On the other hand, specific structural methods would suggest that lyophilization of plasmid DNA would have little if any negative effect. Biofunctional assays showed that lyophilization of plasmid DNA causes a significant decrease in functional activity, showing that in some instances functional assays can be more sensitive than structural assays. Secondly, it was hypothesized that environmental factors will affect the stability of plasmid in solution. This hypothesis was shown to be true, with plasmid DNA being degraded by extremes in temperature and pH. Most interesting was the buffer catalysis effect exerted by citrate. This stresses the importance of a rational design strategy for the formulation of plasmid DNA as a drug product. Specifically, plasmid DNA was exceedingly stable in solution over a wide range of pH and temperature extremes. However, the choice of buffer is of more importance than has typically been considered in the past. Based on the literature review and the data presented here, it is most advisable to formulate plasmid DNA at neutral to slightly basic conditions, using a low buffer concentration that will minimize any potential buffer effects.

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105 The third hypothesis was that the stability of lyophilized plasmid DNA to environmental factors will be higher than plasmid DNA in solution but that the lyophilization process may damage plasmid DNA via conformational strain caused by the removal of the DNA hydration sphere. These data imply that plasmid DNA damage after lyophilization is mediated by some sort of conformational change, as supported by circular dichroism spectra. This conformational change is not prevented by all carbohydrates tested, but all protectants did protect against the hyperchromic effect. This suggests that lyoprotectants may decrease denaturation of the plasmid DNA caused by the lyophilization process. Finally, this project tested the hypothesis that increasing levels of endotoxin contamination would result in reduced functionality of plasmid DNA in tissue culture methods. It was clear that several factors besides the plasmid and delivery vector would influence the activity of non-viral gene delivery systems. Endotoxin contamination could potentially impact transfection efficiency via competition with plasmid DNA for cationic liposome binding, but this would not be expected with typical GLP or GMP preparations used in clinical studies where the endotoxin levels range from 10 1000 EU/mg DNA. At these reduced

PAGE 118

106 endotoxin levels, the competition between endotoxin and DNA for the cationic liposome is at best marginal. An effect could be seen, with increased transfection variability, at SO EU/ml. This level of endotoxin contamination can occur with small scale plasmid preparations used for in vitro cell transfections, and potentially affects the results seen in many in vitro studies. This study has examined many factors that can affect the pharmaceutical formulation and development of plasmid DNA. While there are a number of conditions that can damage plasmid DNA, the rational design and formulation of plasmid DNA as a drug should be possible in the future using the studies presented here.

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107 LIST OF REFERENCES Aida, Y., and Pabst, M. J. (1990). ~Removal of endotoxin from protein solutions by phase separation using Triton X-114." Immunol. Meth. 132: 191-195. Anchordoquy, T. J., Kroll, D. J., and Carpenter, J. F. (1997). ~stabilization of DNA/lipid complexes during freeze-drying." Pharm. Res. 14(11): S641-S642. Bailleal, B., Galiegue-Aouitrina, S., and Loucheux Lefebrue, M. (1984). ~conformations of poly(dG-dC) poly(dG-dC) modified by the O-acetyl derivative of the carcinogen 4-hydroxyamino-quinoline 1-oxide." Nuc. Acids Res. 12(20): 7915-27. Bottoms, G.D. (1982). ~Thromboxane prostaglandin 1-2 epoprostenol and the hemodynamic changes in equine endotoxin shock." Am. J. Vet. Res. 43: 999-1002. Caplen, N., J., Gao, X., Hayes, R., Elaswarapu, R., Fisher, G., Kinrade, E., Chakera, A., Schorr, J., Hughes, B., Dorin, J. R., Porteous, D. J., Alton, E.W. F. W., Geddes, D. M., Coutelle, C., Williamson, L. H., and Gilchrist, C. (1994). ~Gene therapy for cystic fibrosis in humans by liposome-mediated DNA transfer: the production of resources and the regulatory process." Gene Ther. 1: 139-147. Carpenter, J. F., and Crowe, J. H. (1989). ~The mechanism of cryoprotection of proteins by solutes." Cryobiology 25: 244-255. Cotton, M., Baker, A., Saltik, M., Wagner, E., and Buschle, M. (1994). ~Lipopolysaccharide is a frequent contaminant of plasmid DNA preparations and can be toxic to primary human cells in the presence of adenovirus." Gene Ther. 1: 239-246.

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108 Crowe, J. H., Carpenter, J. F., Crowe, L. M., and Achordoquy, T. J. (1990). ~Are freezing and dehydration similar stress vectors? A comparison of modes of interaction and stabilizing solutes with biomolecules." Cryobiology 27: 219-231. Crowe, J. H., Crowe, L. M., Carpenter, J. F., Rudolph, A. S., Wistrom, C. A., Spargo, B. J., and Anchordoguy, T. J. (1988 ) ~Interactions of sugars with membrane." Biochim Biophys Acta 947: 367-384. Durland, R., and Eastman, E. (1998). ~Manufacturing and quality control of plasmid-based gene expression systems." Adv. Drug Deliv. Rev. 30: 33-48. Epstein, J., Lee, M. M., Kelly, C. E., and Donahoe, P. K. (1990). ~Effect of endotoxin on mammalian cell growth and recombinant protein production." In Vitro Cell. Dev. Biol. 26: 1121-1122. F.D.A (1985 ) ~Inspection Technical Guide Number 40: Bacterial Endotoxins/Pyrogens." U.S. Food and Drug Administration. Falk, M., Hartman, K. A., and Lord, R. C. (1963). ~Hydration of deoxyribonucleic acid II: an infrared study." J. Am. Chem. Soc. 85: 387-391. Finnegan, M., Herbert, K., Evans, M., Griffiths, H., and Lunec, J. (1996). ~Evidence for sensitisation of DNA to oxidative damage during isolation." Free Rad. Biol. Med. 20(1): 93-98. Fletcher, J. R., and Ramwell, P. W. prostacyclin on endotoxin shock platelet aggregation in dogs." 308. (1980). ~The effects of and endotoxin-induced Circ. Shock 7(3): 299Frederico, L.A., Kunkel, T. A., and Shaw, B. R. (1990). A sensitive genetic assay for the detection of cytosine deamination: determination of rate constants and the activation energy." Biochemistry 29: 2532-2537. Freshney, R. I. (1994). Culture of Animal Cells 3rd Edition. N~w York, Wiley-Liss Inc. Friedmann, T. ( 1997) Overcoming the obstacles to gene therapy." Sci Am 276(6): 96-101.

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111 Nabel, G. J., Nabel, E. G., Yang, Z., Fox, B. A., Plautz, G. E., Xiang, G., Huang, 1., Shu, S., Gorden, D., and Chang, A. E. (1993). ~Direct gene transfer with DNA liposome complexes in melanoma: expression, biologic activity and lack of toxicity in humans." Proc. Natl. Acad. Sci. USA 90: 11307-11311. New, R.R. C. (1990). Liposomes: a practical approach. New York, IRL Press. Nishimura, Y., Tsuboi, M., Uegi, S., Ohkuba, M., and Iekahara, M. (1985). ~salt induced conformational transition between A and Z forms of r(CGCGCG) as revealed by raman spectrophotometric study." Nuc. Acid Symp. Ser. 16: 25-8. Oka, H., Harada, K., Suzuki, M., Nakazawa, H., and Ito, Y. (1991). ~Foam counter-current chromatography of bacitracin: II continuous removal and concentration of hydrophobic components with nitrogen gas and distilled water free of surfactants or other additives." J. Chromatogr. 538: 213-218. Ozaki, K., and Hayashi, M. (1997). ~The effects of glucose oligomers (maltodextrans) on freeze-drying liposomes." Chem. Pharm. Bull. 45(1): 165-170. Poelstra, K., Bakker, W., Klok, M., and Meij er, D. (1997a) ~A physiologic function for alkaline phosphatase: endotoxin detoxification." Lab. Invest. 76: 319-327. Poelstra, K., Bakker, W., Klok, P., Kamps, J., Hardonk, M., and Meijer, D. (1997b). ~Deposphorylation of endotoxin by alkaline phosphatase in Vivo." Am. J. Path. 151(11631169). Poxon, S. W., Mitchell, P. M., Liang, E., and Hughes, J. A. (1996). ~Dendrimer delivery of oligonucleotides." Drug Delivery 3: 255-261. Rietschel, E. T., Kirikae, T., Schade, U., Ulmer, A. J., Holst, 0., Brade, H., Schmidt, G., Mamat, U., Grimmecke, H., Kusumoto, S., and Zahringer, U. (1993). ~The chemical structure of bacterial endotoxin in relation to bioactivity." Immunobiol. 187: 169-190.

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113 Talsma, H., Cherng, J. Y., Lehrmann, H., Kursa, M., Ogris, M., Bennink, W. E., Cotten, M., and Wagner, E. (1997). ~stabilization of gene delivery systems by freeze drying." Int. J. Pharm. 157(2): 233-238. Tan, Y., Xu, M., Tan, X., Tan, X., Wang, X., Saikawa, Y., Nagahama, T., Sun, X., Lenz, M., and Hoffman, R. M. (1997). ~overexpression and large-scale production of recombinant L-methionine-alpha-deamino-gama mercaptomethane-lyase for novel anticancer therapy." Protein Exp. Purif. 9: 233-245. Tao, N. J., and Lindsay, S. M. (1989). ~structure of DNA hydration shells as studied by Raman spectroscopy." Biopolyrners 28: 1019-1030. Tsuji, K., and Lewis, A. R. (1978). ~Dry-heat destruction of lipopolysaccharide: dry-heat destruction kinetics." Environ. Microbiol. 36: 710-714. Umrania, Y., Nikjoo, H., and Goodfellow, J. M. (1995). ~A knowledge based model for DNA hydration." Int. J. Radiat. Biol. 67: 145-152. Weber, M., Moller, K., Welzeck, M., and Schorr, J. (1995). ~Effects of lipopolysaccharide on transfection efficiency in eukaryotic cells." Biotechniques 19(6 ) : 930-940. Wicks, I. P., Howell, M. L., Hancock, T., Kohsaka, H., Olee, T., and Carson, D. A. (1995). ~Bacterial lipolysaccharide copurifies with plasmid DNA; implications for animal models and human gene therapy." Hurn. Gene Ther. 6: 317-323. Williams, N. A., and Polli, G. P. (1984). ~The lyophilization of pharmaceuticals: a literature review." J Parenter Sci Technol 38(2): 48-59. Wils, P., Escriou, V., Warnery, A., Lacroix, F., Ollivier, M., Crouzet, J., Mayaux, J. F., and Scherman, D. (1997). ~Efficient purification of plasmid DNA for gene transfer using triple-helix affinity chromatography." Gene Ther. 4: 323-330. Wong, D., and Parasrampuria, J. (1997). ~Pharmaceutical excipients for the stabilization of proteins." Pharm. Tech. 21 (10) : 34-50.

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114 Xing, Z., Jordana, M., Kirpalani, H., Driscoll, K. E., Schall, T. J., and Gauldie, J. (1994). ~cytokine expression by neutrophils and macrophages in vivo: endotoxin induces tumor necrosis factor alpha, macrophage inflammatory protein-2, interleukin-1 beta, and interleukin-6 but not RANTES or transforming growth factor-beta 1 rn.RNA expression in acute lung inflammation." Am. J. Respir. Cell Mol. Biol. 10: 148153. Xu, Y., and Szoka, F. (1996). ~Mechanism of DNA release from cationic liposome/DNA complexes used in cell transfection." Biochem 35(18): 5616-5623.

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BIOGRAPHICAL SKETCH Scott William Poxon was born on January 4, 1971, in Ridgewood, NJ. He lived in New Jersey for his first 7 years, whereupon his family moved to Whitehall, New York. After residing in New York for 3 years, Scott moved to Sarasota, FL. Scott finished elementary school and was admitted to Pine View School for the Gifted, which he attended through the 12 th grade. While at Pine View, Scott was named a National Merit Semi-Finalist and was a participant at the 1987 Florida State University Summer Science and Math Program, where he was first introduced to molecular biology. Scott matriculated from Pine View in 1989, with honors, and subsequently attended the University of Central Florida. Scott was a member of Omicron Delta Kappa, Phi Mu Alpha, and Sigma Chemical Company Chi while at Central Florida. He received a bachelors degree in microbiology and molecular biology and a minor in violin performance in 1994. Scott then attended the University of Florida, where he completed a Ph.D. in pharmaceutics and interdisciplinary toxicology. While at the University of Florida, Scott was 115

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116 a member of the American Association of Pharmaceutical Scientists. He also was the Graduate Research Association of Students in Pharmaceutics representative for the University of Florida, and was the organization's 1997 chair, hosting the annual meeting at the University of Florida. He served terms as Secretary of the College of Pharmacy Graduate Student Council, and Departmental Representative to the University of Florida Graduate Student Council. Scott received several awards while at the university including the College of Pharmacy Annual Oral Competition, the Liberty Fellowship, the Advanced Predoctoral Fellowship in Pharmaceutics from the Pharmaceutical Research and Manufacturers of America Foundation, and the Biotechnology Grant from the Parenteral Drug Association Foundation for Pharmaceutical Sciences.

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Hughes, Chair sistant Professor of Pharmaceutics I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Associate Professor of Pharmaceutics I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Edfk;;r ~ Associate Professor of Pharmacology and Therapeutics I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Phil sophy.

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Ian Tebbett Professor of Medicinal Chemistry This dissertation was submitted to the Graduate Faculty of the College of Pharmacy and to the Graduate School and was accepted as par requirements for the degree o May 1999 I Dean, Graduate School

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UNIVERSITY OF FLORIDA 1111111111111111I IIIII IIIII II IIIIII IIII IIII Ill llll Ill llll lllll I I 3 1262 08555 2973


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