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The effect of DNA supercoiling on transcription initiation at the glnAp2 promoter

Honors Thesis

Presented to

Brandeis University Biological Physics Program

Jeff Gelles, Advisor

In Partial Fulfillment of the Requirements for the Degree

Bachelors of Science

in Biological Physics

by

Abby Knecht

May 2015

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Copyright by

Abigail Knecht

© 2015

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Abstract: A quantitative kinetic mechanism for transcription initiation at the bacterial

promoter glnAp2 was previously determined using single molecule experiments

with relaxed DNA. DNA in E. Coli cells, however, is rarely relaxed but is instead

negatively supercoiled. Bulk experiments in the literature report a 10 to 60 times

increase in the overall rate of transcription initiation at the glnAp2 promoter with

negatively supercoiled DNA compared to relaxed DNA. To locate where in the

mechanism this increase in rate occurs, I created circular DNA that contained the

glnAp2 promoter, biotin for attachment to a microscope slide, and a dye for

visualization. Initiation was then measured using single molecule techniques on

both nicked and negatively supercoiled forms of this template. The overall rate of

transcription initiation was seven times faster on negatively supercoiled DNA

compared to on nicked DNA. The rate of RNA polymerase binding, however, was

only 1.5 times faster on supercoiled DNA, and the rate of open complex decay on the

supercoiled and nicked templates was not significantly different. Initial experiments

on the RNA polymerase closed complexes suggest that their stability on negatively

supercoiled and nicked templates is not very different. Based on those results, I

hypothesize that isomerization from closed to open complex is most likely the step

in the mechanism that accounts for the difference in overall rate of transcription

initiation between relaxed and supercoiled DNA.

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Table of Contents:

Chapter 1 – Introduction

Supercoiling Theory

Transcription Initiation

TIRF Microscopy

Supercoiling and Transcription Initiation

Chapter 2 – Synthesis of Circular Transcription Templates

Results

Digesting pJES534

Assembling Dye/Biotin Fragment

First Ligation

Determining the Circular Template

Nicking and purification of the circular template

Second Ligation and Gyrase Reaction

No promoter Circular Templates

Calculations

Materials and Methods

DNA plasmids and oligonucleotides

Enzymes

Labeling the oligonucleotide

Chapter 3 – Transcription Initiation Mechanism

Results

Specificity of Initiation

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Overall rate of initiation

Rate of first binding of RNAP

Closed Complexes

Open complex dissociation


TIRF Microscope and Slide Preparation

Solutions

Fit Equations

Chapter 4 – Discussion

References

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List of Figures

Chapter 1 – Introduction

Figure 1.1: DNA Supercoiling – Linking Number, Twist, and Writhe

Figure 1.2: TIRF Microscope

Chapter 2 – Synthesis of the Circular Transcription Templates

Figure 2.1: Overall procedure for constructing the Nicked and Supercoiled

Templates

Figure 2.2: Restriction Fragment

Figure 2.3: Labeling and Annealing the Dye/Biotin Fragment

Figure 2.4: First Ligation

Figure 2.5: Testing for the circular template with Gyrase and Nt.AlwI

Figure 2.6: Nicking Digest and Gyrase Reaction

Figure 2.7: Circular Templates with and without a promoter

Chapter 3 – Transcription Initiation Mechanism

Figure 3.1: Testing transcription initiation using the circular templates

Figure 3.2: Tracing intensity of labeled probe

Figure 3.3: Transcription initiation fraction on different circular templates.

Figure 3.4: Rates of transcription initiation and 𝜎!"-‐RNAP binding

Figure 3.5: Closed Complex Dwell Times

Figure 3.6: Dissociation of the Open Complex


Figure 4.1: Mechanism of Transcription Initiation for 𝜎!"-‐RNAP on Circular

DNA

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Chapter 1: Introduction

Supercoiling Theory

DNA is commonly portrayed as a linear double helix, but DNA is not linear

and relaxed in the cell. DNA in cells is supercoiled, either in bacterial plasmids, or in

chromosomal DNA both in bacteria and eukaryotes (Mirikin 2002). Supercoiling

occurs when the DNA double helix is either tightened (known as positive

supercoiling) or loosened (known as negative supercoiling). In cells, chromosomal

DNA is negatively supercoiled both so it can fit into the cell and so that proteins can

more easily bind to the DNA and dehybridize the hydrogen bonds between base

pairs (Watson 2007).

Supercoiling can mathematically be described with three variables: linking

number (Lk), twist (Tw), and writhe (Wr). When a closed circular piece of DNA is

flattened, as it is in Figure 1.1A, the linking number is defined by the number of

times one strand of the DNA crosses over the other strand (Mirkin 2002). As such,

the linking number is always an integer. The twist is defined as the total number of

helical turns in the circular DNA. The writhe describes the 3D shape of the DNA. It

is defined by the number of times the long axis of the DNA crosses over itself

(Watson 2007). Writhe presents itself as either a plectoneme (Figure 1.1B, the

figure eight shape in) or a toroid and can be either greater than, less than, or equal

to zero (Marko and Neukirch 2012). These quantities are related by the simple

equation:

𝐿𝑘 = 𝑇𝑤 +𝑊𝑟 (1.1)

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A relaxed DNA wraps around its axis about once every 10 base pairs.

Therefore, the linking number of a relaxed DNA (LkO) can be approximated by

dividing the number of base pairs by 10. When the DNA is relaxed, the writhe of the

molecule is zero, so 𝐿𝑘 = 𝐿𝑘! = 𝑇𝑤 (Figure 1.1A). Supercoiled DNA is defined by

DNA that has a linking number does not equal its LkO. Mathematically, the amount

of supercoiling can be described by the difference between the current linking

number of the DNA and its LkO (equation 1.2).

Δ𝐿𝑘 = 𝐿𝑘 − 𝐿𝑘! (1.2)

Since the linking number depends on the number of base pairs in the DNA,

the superhelical density (𝜎) is most commonly used to measure the amount of

supercoiling. The superhelical density is defined by equation 1.3:

𝜎 = !!"!"! (1.3)

When the superhelical density is greater than zero the DNA is positively

supercoiled, when it is less than zero the DNA is negatively supercoiled, and when it

is equal to zero the DNA is relaxed.

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Figure 1.1: DNA Supercoiling – Linking Number, Twist, and Writhe. A. Example of a relaxed circular plasmid where the linking number is equal to the twist, so the writhe is zero. B. The same molecule as A, but a topoisomerase has decreased the linking number by 4. The change in linking number created a writhe of -‐4 in the molecule, while the twist remains the same. This molecule is supercoiled. C. The DNA here has the same linking number as B, but here the writhe is 0 and the twist has changed. The change in twist but not writhe causes the base pairs to become disrupted. Figure from Watson 2007. Cells are able to manipulate the superhelical density of their DNA by using

topoisomerases (Figure 1.1 A-‐B). Topoisomerases are enzymes that change the

linking number of a DNA by cutting either one strand (type I topoisomerases) or

two strands (type II topoisomerases) and then re-‐ligating them with a changed

linking number. Most topoisomerases simply relax supercoiled DNA back to its

relaxed state, but some, such as the E. Coli enzyme gyrase, can induce negative

supercoils into DNA in the presence of ATP (Bates and Maxwell 1989; Gellert et al.

1976). I used gyrase in the experiment to induce negative supercoils into the

circular template.

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Transcription Initiation

Transcription is the process by which a gene is copied by the enzyme RNA

polymerase (RNAP) into a single strand of RNA. Transcription consists of three

main parts: initiation, elongation, and termination (Weaver 2012). Transcription

initiation is where an RNAP finds a certain sequence in the DNA before the

beginning of the gene, known as the promoter, and opens the DNA to form an open

complex. Elongation occurs just after initiation when the RNAP creates the RNA by

hybridizing nucleotides to the DNA. When the RNA polymerase has finished

transcribing the gene, it dissociates from the DNA in a process known as

termination.

Here I studied transcription initiation in vitro using proteins from E. Coli. The

RNA polymerase holoenzyme in bacteria consists of the RNA polymerase core and a

specificity subunit called a sigma (𝜎) factor. Depending on the promoter involved, a

specific 𝜎 factor is needed for the RNAP to begin transcribing. The 𝜎!" factor that I

study here is responsible for activating transcription on genes that are needed for

Nitrogen metabolism in E. Coli as well as other functions (Mauri and Klumpp 2014,

Weaver 2012). Unlike some other sigma factors in E. Coli, the 𝜎!" RNAP

holoenzyme is unable to initiate transcription on its own and is in need of an

activator (Wedel and Kustu 1995). The well-‐established activator for the glnAp2

promoter is NtrC (nitrogen regulating protein C). NtrC hydrolyzes ATP or GTP to

couple the energy involved in hydrolyzation to open the DNA at the promoter

forming the open complex (Wedel and Kustu 1995).

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TIRF Microscopy

Transcription is a very dynamic process with proteins binding quickly and

then dissociating. While bulk experiments can be elusive in determining overall

rates, single molecule experiments are more useful in determining the

heterogeneity of the dynamic process. The single molecule experiments I use here

are with total internal reflection fluorescence (TIRF) microscopy. TIRF microscopy

reduces background fluorescence by only exciting approximately 100-‐nm above the

surface of the glass slide (Friedman, Chung, and Gelles 2006). The TIRF microscope

used in my project can simultaneously excite the sample at multiple wavelengths by

spatially separating the excitation and emission beams (Figure 1.2, Friedman,

Chung, and Gelles 2006). With the ability to simultaneously excite multiple

wavelengths, the TIRF microscope can be used to study single molecules interacting

using CoSMoS (Co-‐localization single-‐molecule spectroscopy) (Friedman and Gelles

2012).

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Figure 1.2: TIRF Microscope. Diagram of the TIRF Microscope from Friedman, Chung, and Gelles 2006. The excitation beam (green arrows, 𝜆!) is reflected by the dichroic mirror and then is totally internally reflected at the boundary between the aqueous buffer and the glass slide. The longer emission beam (red arrows, 𝜆!) is transmitted through the dichroic mirror.

Supercoiling and Transcription Initiation

Transcription initiation at the glnAp2 promoter is a dynamic multi-‐step

process. The entire mechanism of transcription initiation at this promoter was

determined using CoSMoS with linear DNA attached to the slide (Friedman and

Gelles 2012). The mechanism includes two closed complexes followed by the

formation of the open complex. After the open complex, the transcription elongation

complex (TEC) forms and transcription elongation begins as long as NTPs are

present.

While the mechanism of transcription initiation was previously determined

using linear DNA, DNA in cells is negatively supercoiled. The stability of the open

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complex for the 𝜎!" RNAP changes depending on whether it forms on negatively

versus positively supercoiled DNA (Revyakin, Ebright, and Strick 2004). There is

also evidence that the formation of the 𝜎!" RNAP open complex peaks at a

superhelical density of -‐0.024, while the 𝜎!" RNAP peaks at -‐0.067, showing that

transcription initiation depends on supercoiling for both RNA polymerases

(Whitehall, Austin, and Dixon 1992). Bulk experiments similarly determined that

the overall rate of initiation at the glnAp2 promoter is 10 to 60 times faster on

negatively supercoiled DNA compared to relaxed DNA (Schulz, Langowski, and

Rippe 2000). However they give no indication of where this increase in rate occurs

in the multi-‐step process of transcription initiation.

The goal of my experiments was to determine the mechanism of

transcription initiation on supercoiled DNA at the glnAp2 promoter. There were

two main parts of my project. I first created a circular, supercoiled piece of DNA

that could be attached the microscope slide using biotin, had a fluorescent dye

attached, and had the sequences necessary for transcription initiation and

elongation. After the circular template was completed, I performed single molecule

TIRF microscope experiments to determine the mechanism of transcription

initiation using the supercoiled DNA.

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Chapter 2: Synthesis of Circular Transcription Templates

Results

In order to perform TIRF experiments to determine where in the mechanism

supercoiled DNA transcribed faster, I had to first construct a supercoiled circular

template. The template needed to have the necessary sequences for transcription

initiation and elongation for a 𝜎!" RNAP, a fluorescent dye to be visualized in the

microscope, and a biotin moiety to attach the template to a streptavidin-‐coated

microscope slide. The construction of the circular template was similar to that of

Kalkbrenner et al. (Kalkbrenner, Arnold, and Tans 2009). In summary (Figure 2.1),

the sequences for transcription initiation were isolated as a restriction fragment of a

stock plasmid and then ligated with a synthesized oligonucleotide containing the

biotin and fluorescent dye (dye/biotin fragment). The correct circular template

was then purified from the ligation reaction. After purification, the circular template

was supercoiled using the enzyme gyrase.

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Figure 2.1: Overall procedure for constructing the Nicked and Supercoiled Templates. The restriction fragment (760 base pairs) is in green while the dye/biotin fragment (32 base pairs) is in purple. The E and H correspond to the sticky ends that resulted from the restriction digest by EcoRI and HindIII respectively. The final circular templates are highlighted in red surrounded by black boxes.

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Digesting pJES534

The sequences needed for transcription initiation and elongation (the

restriction fragment) were purified from the plasmid pJES534 (Figure 2.1A,

Friedman and Gelles 2012) and consisted of three main parts: the promoter

sequence, the enhancer sequences, and the repeating cassette sequence. The

plasmid was digested with EcoRI and HindIII, each having a binding site on either

end of the restriction fragment (Figure 2.2A). The restriction fragment was then gel

purified to isolate it from the rest of the plasmid. The promoter sequence I used for

this experiment is the glnAp2 promoter, which requires the 𝜎!" initiation factor and

the activator NtrC to activate transcription initiation (Vidangos et al. 2013; Wedel

and Kustu 1995). The enhancer for NtrC consists of two binding sites upstream of

the promoter (Figure 2.2B). Directly downstream of the promoter are the repeating

cassettes that will be transcribed by the RNA polymerase (Figure 2.2 B, C). Due to

the construction of the repeating cassette sequence, the RNA transcribed will not

contain any uracil within the repeats. If UTP is not added to the reaction, the

polymerase will stall when it reaches a thymine base at the end of the repeating

cassettes.

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Figure 2.2 Restriction Fragment. A. Plasmid pJES534 with the EcoRI and HindIII restriction sites labeled. The green segment is the restriction fragment that is gel purified. B. The restriction fragment after it is purified. The locations of the enhancer sequences, promoter sequence, and repeating cassette sequence are shown in their respective locations. C. The sequence of a single cassette. The repeating cassettes consisted only of A, C, and G NTPs.

Assembling Dye/Biotin Fragment

To create the dye/biotin fragment, two different synthetic oligonucleotides

were annealed together (Figure 2.3A). Initially, one of the oligonucleotides

contained biotin and the other that had an amine attached. Since I wanted to create

a circular template, the biotin and the fluorescent dye had to be attached in the

middle of the oligonucleotide, as attaching it at the ends would make it impossible to

close the circle. In addition, I was worried that kinks in the circular template from a

missing base that would interfere with the supercoiling of the circular templates.

Instead the oligonucleotides were ordered with the dye and biotin attached to a

thymine base so that none of the base pairs would be disrupted in the final circular

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template (Figure 2.3 B-‐C). The amine group was replaced with an Alexa Fluor 488

fluorescent dye using an amine-‐ester reaction. After the dye was attached, the two

oligonucleotides were annealed and then phosphorylated with a kinase to form the

dye/biotin fragment (Figure 2.3D).

Figure 2.3: Labeling and Annealing the Dye/Biotin Fragment. A. Schematic procedure to assemble the dye/biotin fragment. The purple fragments represent the synthesized oligonucleotides. B. Modified chemical structure of the amine attached to a thymine nucleotide (original structure from IDT). C. Modified chemical structure of the biotin attached to a thymine nucleotide (original structure from IDT). D. Sequence of the dye/biotin fragment after labeling with the Alexa Fluor 488 dye and annealing the two strands together. The red T’s represent the modified thymine bases in B and C. E and H correspond to the 5’ overhangs that result from an EcoRI and HindIII digest respectively (A, D).

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First Ligation

To create the circular template, one restriction fragment had to be attached

to one dye/biotin fragment. The dye/biotin fragment had complementary sticky

ends to the restriction fragment, which facilitated ligation of the two pieces (Figure

2.4A). As expected, the ligation did not only produce the circular template with one

restriction fragment and one dye/biotin fragment. Instead, the ligation produced

many products (some possible products are depicted in Figure 2.4B-‐G). Figure 2.4H

shows the products of the first ligation.

Figure 2.4: First Ligation. A. Cartoon of the two reactants in the first ligation. The restriction fragment (in green) and the dye/biotin fragment (in purple) were ligated together in a 1:1.5 ratio. B -‐ G. Products that could have been formed during the first ligation. Products B-‐D contain two restriction fragments. I hypothesize that they should run in the 1.5-‐2 kb range on an agarose gel. Products E-‐G contain only one restriction fragment. Theoretically they should run around the 1 kb range on an agarose gel. H. Agarose gel electrophoresis of the first ligation (imaged by Alexa Fluor 488 fluorescence). Bands 1-‐5 were excised for subsequent use. The numbers to the left of the gel represent the positions of the 1kb ladder for this gel.

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Determining the Circular Template

By utilizing how topoisomers run on an agarose gel, the identity of the bands

in Figure 2.4H could be determined. On an agarose gel, an open circle, a linear DNA,

and a highly supercoiled circle of the same length will run differently. A nicked or

open circle runs the slowest, while a highly supercoiled circle will run the fastest.

The linear DNA as well as the other topoisomers of the same length will run

somewhere between the open circle and the highly supercoiled circle (Keller 1975).

As such, the circular template (Figure 2.4F) and its topoisomers (Figure 2.4G)

should run close to the position of the linear template (Figure 2.4E) on the gel.

In Figure 2.4 H there are 4 bands that are close to the position of the linear

template on the gel (at 800 base pairs). To determine if these were topoisomers of

the circular template (Figure 2.4G) or whether they were other linear templates

(Figure 2.4E), each band was gel extracted (Figure 2.4H, bands 1-‐4) as well as the

next highest in the gel for a control (Figure 2.4H, band 5). The extracted DNA was

tested with gyrase and the nicking enzyme Nt.AlwI to see whether they were

topoisomers of the circular template (Figure 2.4G) or linear templates (2.4E). If they

were all different linear templates, then neither the gyrase nor the nicking enzyme

would have an effect on the migration of the bands. If they were topoisomers of the

circular template on the other hand, then they would all shift to the same position

with the addition of either gyrase or Nt.AlwI.

When incubated with gyrase, bands 2-‐4 migrated to the same position

(Figure 2.5A, lanes d, f, and h). This is consistent with the hypothesis that bands 2-‐4

are topoisomers of the circular template. The control, band 5, however did not

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migrate to the same position as the others when it was supercoiled (Figure 2.5A,

lanes i-‐j). Since it ran closer to the 2kb range, I concluded that it was likely a linear

dimer (Figure 2.4B). From previous experiments I determined that band 1 was a

linear template because it consistently ran at 800 base pairs, which is consistent

with where the linear template would run.

Figure 2.5: Testing for the circular template with Gyrase and Nt.AlwI. A. Agarose gel of the gyrase experiment. The numbers for DNA refer to the gel extracted bands from Figure 4H. The (+)-‐sign indicates that the sample was incubated with gyrase at 37˚C to supercoil the DNA. Lane a is a 1kb ladder. B. Agarose gel of the nicking digest using Nt.AlwI. The (+)-‐sign indicates that the sample was incubated with the nicking enzyme Nt.AlwI. Both gels were incubated with ethidium bromide after the gel was run and then imaged for ethidium bromide. C. Models of the 5 bands based on the gyrase and nicking digest experiments. Band 1 is the linear template, bands 2 and 3 are topoisomers of the circular template, band 4 is the relaxed circular template, and band 5 is a linear dimer.

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To confirm that bands 2-‐4 were all topoisomers of the same circular

template, I did the same experiment but incubated the extracts with the nicking

enzyme Nt.AlwI (Figure 2.5B). After the incubation with Nt.AlwI, bands 2-‐4

migrated to the same position on the agarose gel (Figure 2.5B, lanes b, d, and f).

These results confirm those from the gyrase experiment, that bands 2-‐4 are all

topoisomers of the circular template. Just as it did in the gyrase experiment, there

was no difference in the migration of band 5 with or without the nicking enzyme; it

still ran at 1.5kb (Figure 2.5B, lane h). The gyrase supercoiling experiment

combined with the nicking digest confirms that the four products near 1 kb in the

first ligation are topoisomers of the circular template (Figure 2.4G). Figure 2.5 C

gives a summary of what each of the five bands are based on the gyrase and nicking

enzyme tests.

Nicking and purification of the circular template

The next step was to purify the circular template from the first ligation

reaction. To do so, the first ligation was nicked using the Nt.AlwI enzyme and run on

a gel (Figure 2.6A, lane: First Ligation Nicked), and then the relaxed circular

template was gel purified (Figure 2.6A, lane: Nicked Circular Template). Some of the

purified nicked circular template (Figure 2.6B) was stored for a comparison to the

supercoiled DNA during the TIRF experiments, while the rest was used to make the

supercoiled circular template.

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Figure 2.6: Nicking Digest and Gyrase Reaction. A. Agarose gel that illustrates the procedure to isolate the nicked and supercoiled circular templates. The gel electrophoresis was run without ethidium bromide, but was incubated with ethidium bromide after the gel was finished running and then imaged for ethidium bromide. B and C. Cartoon diagrams of nicked and supercoiled circular templates respectively. Green represents the promoter fragment while purple is the dye/oligo fragment. Second Ligation and Gyrase Reaction:

With the nicked circular template purified, the next step was to negatively

supercoil it (Figure 2.1). Since gyrase is unable to supercoil DNA that is nicked, the

nicked circular template needs to incubated with ligase before it can be supercoiled.

The second ligation resulted in two distinct products (Figure 2.6A, lane: Second

Ligation). The probability that a circular piece of DNA has a certain superhelical

density without the aid of topoisomerases is given by equation 2.1 and 2.2. With

these equations, I calculated the probability that the circular template was relaxed

to be 66% and the circular template to have a change in linking number ±1 to be

33%. The ratio in the intensities of bands a and b in Figure 2.6A was approximately

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50:50. Despite the difference between theoretical ratio and the measured one, it is

probable that band a is the relaxed circular template and band b is the topoisomers

with a ±1 change in the linking number. Once the nicks had been removed from the

circular template by the second ligation, the circular template was incubated with

gyrase to form a negatively supercoiled circular template (Figure 2.6A, lane:

Supercoiled circular template, and 2.6C).

No Promoter Circular Templates

A circular template without a promoter was also prepared using the method

described above (Figure 2.1). The restriction fragment for the no promoter circular

template was isolated from a plasmid that was the same as pJES534, but it had the

glnAp2 promoter sequence (-‐1 to -‐27 with respect to the transcription start site)

removed. Otherwise, all the reactants that were used to assemble the circular

template with a promoter were used for the synthesis of the circular template

without the promoter. Figure 2.7 shows the final circular template both with the

glnAp2 promoter (A) and without the promoter (B). It also describes the locations

of the sites where Nt.AlwI nicks the DNA and the restriction digest sites for EcoRI

and HindIII.

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Figure 2.7: Circular Templates with and without a promoter. A-‐B. Diagrams of the final circular template with (A) and without (B) a promoter. The renderings describe the locations of the sequences needed for transcription initiation and elongation, as well as the positions of the restriction sites, biotin, and dye positions. The nicking sites do not interfere with any of the sequences needed for transcription initiation and elongation. Calculations

To find the probability that the circular template is in a particular energy

state without assistance from an outside enzyme, I used the Boltzmann distribution

𝑃(𝐸!) =!!!

!!! !!!

!(!) (2.1)

where Δ𝐺 is the change in the Gibbs free energy of a particular state from its relaxed

state, gi is the number of times that energy state can occur, T is the temperature, kB

is the Boltzmann constant, and z(t) is the partition function. To calculate Δ𝐺 for one

mole of topoisomer, I used the equation

Δ𝐺 = 10𝑅𝑇𝑁𝜎! (2.2)

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where R is the gas constant, N is the number of base pairs, and 𝜎 is the superhelical

density (Seidl and Hinz 1984). The relaxed state occurs when the superhelical

density is zero and the circular DNA is relaxed. As explained in equation 1.3, 𝜎 is

defined by Δ𝐿𝑘 𝐿𝑘!. In my experiment, the circular template is 792 base pairs (N).

For a relaxed topoisomer (Δ𝐿𝑘 = 0), Δ𝐺 = 0, and the probability that it will

occur is 66%. For a topoisomer where Δ𝐿𝑘 = ±1, Δ𝐺 = 𝑒!!.!" ≈ 0.25. In this case

since both the positive and negative supercoiled topoisomers have the same energy,

gi = 2. Therefore, the probability that the topoisomer will be supercoiled by one

turn in either the positive or negative direction is 33%. Applying the same principle

but with a Δ𝐿𝑘 = ±2, the probability is only 0.56%. Therefore it is unlikely that a

Δ𝐿𝑘 = ±2 or higher would occur without assistance from an outside source (such as

a topoisomerase).


DNA plasmids and oligonucleotides

The plasmids used to isolate the restriction fragments were pJES534 that

either contained the glnAp2 promoter or did not contain the promoter (Friedman

and Gelles 2012). The plasmids were purified using a Qiagen miniprep kit as per the

manufacturer’s instructions. The oligonucleotides were purchased from IDT with

the biotin and amine already attached as previously described (Figure 2.3 B, C).

Enzymes

The restriction endonuclease HindIII, the nicking enzyme Nt.AlwI, T4 DNA

ligase, and DNA gyrase (E. Coli) were all bought from New England Biolabs (NEB),

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while the restriction endonuclease EcoRI was bought from Promega. During the

digest of the plasmid pJES534, EcoRI and HindIII were incubated together in the

NEB Buffer 2 at 37˚C for 1 hour as per NEB instructions. The ligation reactions were

performed using T4 DNA ligase and the DNA ligase buffer provided per the

manufacturer’s instructions. The nicking reactions using Nt.AlwI were performed

using NEB’s cutsmart buffer and the instructions provided by NEB. The gyrase

reactions were executed per the manufacturer’s instructions using the gyrase buffer

provided.

Labeling the oligonucleotide

The dye used to label the oligonucleotide was an Alexa Fluor 488 carboxylic

acid, 2,3,5,6-‐tetrafluorophenyl ester ~*5-‐isomer, and was purchased from Life

Technologies. The procedure to label was adapted from Life Technologies’s “Amine-‐

Reactive Probe Labeling Protocol”. The reaction buffer used had a final

concentration of 0.33 g/L of amine labeled oligonucleotide, 1.2 g/L of dye, and 0.066

M sodium tetraborate buffer. The reaction was mixed per the manufacturer’s

instructions for the first 2 hours and then left to sit overnight at room temperature.

An ethanol precipitation was performed the next day to remove some of the excess

free dye.

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Chapter 3 – Transcription Initiation Mechanism Results

With the purification of the nicked and supercoiled circular templates, I next

measured the mechanism of transcription initiation at the glnAp2 promoter using

single molecule TIRF microscopy. From the literature, I know that the process of

transcription initiation at this promoter on linear DNA (Friedman and Gelles 2012)

is

𝑅 + 𝑃 ⇄ 𝑅𝑃!! ⇄ 𝑅𝑃!! ⇄ 𝑅𝑃! ⟶ 𝑇𝐸𝐶 + 𝜎!"

where R is the DNA and P is the RNA polymerase. In the mechanism, there are two

closed complexes (𝑅𝑃!! and 𝑅𝑃!!) before the formation of the open complex (𝑅𝑃!).

Compared to the first three forward rate constants, the rate constant from open

complex to the transcription elongation complex (TEC) is much faster. Here I

determined where in the mechanism transcription initiation on the supercoiled

template transcribes faster than on the nicked template.

Specificity of Initiation

To make sure that the circular templates could be used in place of linear

DNA, I tested transcription initiation using the supercoiled and nicked circular

templates. The circular DNA templates were attached to the microscope slide and

then imaged with a blue laser to determine their locations (Figure 3.1 A, B). The

open complex-‐forming solution, consisting of the cy3-‐𝜎!" RNAP holoenzyme, NtrC,

and ATP, was then added to the slide (Figure 3.1 A). I allowed open complexes to

form for 45 minutes, at the end of which the locations of the cy3-‐𝜎!" RNA

polymerases were imaged with a green laser (Figure 3.1C). Finally I added the ATP,

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CTP, and GTP as well as the cy5 probe complementary to a single cassette sequence

to begin elongation. A red laser was then used to visualize where elongation was

occurring (Figure 3.1D). The circular templates, the cy3-‐𝜎!" RNAP, and the cy5-‐

probe could all be seen on the microscope and I could see co-‐localization of the

RNAP and probe to the circular templates (Figure 3.1B-‐D).

Figure 3.1: Testing transcription initiation using the circular templates. A. Diagram of the experimental procedure. B-‐D. Fluorescence images from the TIRF microscope. A blue laser excited the Alexa Fluor 488 dyes on the DNA (B). After the addition of the RNAP, a green laser excited the cy3 dyes on 𝜎!" (C). Ten minutes after the addition of the NTPs and cy5-‐probe, the field was excited with a red laser to see the locations of the probes (D). The arrows mark corresponding locations in the three images (B-‐D).

I next needed to confirm that transcription initiation and elongation were

occurring when the probe co-‐localized with the DNA. To do so, I visualized cy5-‐

probe and cy3-‐𝜎!" binding over 10 minutes at one frame per second after the

addition of NTPs. The longer transcription elongation continued, the more probes

were able to bind to the mRNA, increasing the probe fluorescence intensity (Figure

3.2A). In accordance with the literature, 𝜎!" leaves the RNAP holoenzyme after the

29

formation of the open complex, allowing the transcription elongation complex to

form and progress (Figure 3.2 B; Friedman and Gelles 2012). Therefore

transcription initiation and elongation were occurring as predicted using the

circular templates attached to the slide.

Figure 3.2: Tracing intensity of labeled probe. A. Intensity traces from a single nicked circular template location. The red line represents the intensity over time of the cy5-‐probe while the green line is the intensity of the cy3-‐𝜎!". The measurement started after the 45-‐minute incubation to form open complexes, but just before the addition of NTPs. B. Image gallery of the intensity at the DNA location in the time interval (65s to 95s).

I next wanted to determine how the fraction that initiated changed

depending on which circular template was used. There were four different

constructs that were compared over two days: the supercoiled circular template

with a promoter (Figure 3.3A), the nicked circular templates with and without a

promoter (Figure 3.3 B and C respectively), and the areas without a visible template

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(off DNA) (Figure 3.3D). The fraction that initiated was calculated as the number of

DNA locations that had a co-‐localized cy5-‐probe after 10 minutes divided by the

total number of DNA locations for that particular template. The off DNA locations

were used to measure the amount of nonspecific binding to the surface of the slide

(the background signal). As expected, the fraction that initiated at the nicked and

supercoiled circular templates that contained the promoter (Figure 3.3 A, B) was

significantly higher than the background signal (Figure 3.3D), indicating that

transcription did occur on those circular templates. The nicked template without a

promoter (Figure 3.3C), however, had an initiation fraction comparable to the

background signal (Figure 3.3D), suggesting that it did not initiate. Therefore,

transcription initiation only occurred on circular templates that contained the

promoter, and could occur on both the supercoiled and nicked templates.

The cause of the discrepancy in the fraction that initiated on the nicked

templates on the two days is unclear. It is possible that the difference was caused by

the fact that the first day had CTP and GTP concentrations of 0.5mM each, while the

second day they had concentrations of 1mM each. The higher concentration of CTP

and GTP on the second day may have allowed for faster elongation and therefore an

increased probability that a probe would bind by the end of the ten minutes.

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Figure 3.3: Transcription initiation fraction on different circular templates. Fraction of DNA locations that showed initiation (i.e., a cy5-‐probe co-‐localized) after 10 minutes. The different colors of the bars indicate that the experiments were performed on different days. A. Fraction with transcription initiation on the supercoiled template with a promoter (SC+: n=301). B. Fraction of templates with transcription initiation on the nicked circular template with a promoter (N+ blue: n=229, N+ red: n=415). C. Fraction templates with transcription initiation on the nicked circular template without a promoter (N-‐: n=162). D. Fraction of off DNA sites with a co-‐localized cy5 probe. (Off blue: n=338, Off red: n=352). The error bars indicate standard errors. Overall rate of initiation

Next I determined whether transcription initiation with the supercoiled

circular template was faster than with the nicked circular template. The nicked and

supercoiled circular templates were attached to the slide and incubated with a

solution containing RNAP holoenzyme, NtrC, NTPs, and cy5-‐probe (Figure 3.4A),

allowing steady state initiation to occur in a single reaction mixture. I imaged the

reaction for an hour at two frames per second. To measure the overall rate of

initiation, (𝑅 + 𝑃⟶ 𝑇𝐸𝐶), the cumulative fraction of templates that initiated,

measured as the co-‐localization of the cy5-‐probe with a template for at least 10

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seconds, was plotted against time (Figure 3.4B). By fitting the resulting curves, the

rate of transcription initiation on supercoiled templates was measured to be

2.9± 0.7 ×10!! 𝑠!!, whereas at nicked templates it was measured to be 0.44±

0.18 ×10!!𝑠!!. This gives a seven-‐fold increase in the overall rate of transcription

initiation on the supercoiled template, which agrees with previous results at the

glnAp2 promoter (Schulz, Langowski, and Rippe 2000).

Figure 3.4: Rates of transcription initiation and 𝝈𝟓𝟒-‐RNAP binding. A. Diagram of protocol for TIRF experiment. B. Cumulative plot of the overall rate of transcription initiation. C. Cumulative plot of the time to first RNAP binding. In B and C, the red circles indicate co-‐localization on negatively supercoiled templates (n=233), the blue circles indicate co-‐localization on nicked templates (n=283), while the black circles are the off-‐DNA control locations used to measure the background signal (n=804). The green lines are fits to the data (equation 3.1). Rate of first binding of RNAP

Since on the supercoiled templates, there was faster initiation than on the

nicked templates and these results quantitatively agrees with previous data, this

system can be used to further study the individual rates between steps in the

33

mechanism of transcription initiation. The first step in the mechanism I studied was

the rate of binding of the RNAP to the template, (𝑅 + 𝑃⟶ 𝑅𝑃!!). Using data from

the initiation and elongation experiments (Figure 3.4A), I made a cumulative plot of

the fraction of circular templates with a co-‐localized cy3-‐𝜎!" RNAP versus the time

of its first binding (Figure 3.4C). The rate constant for RNAP binding to the

supercoiled template was measured to be 3.0± 0.4 ×10!𝑀!!𝑠!! and the rate

constant for the RNAP binding to the nicked template was 2.1± 0.3 ×10!𝑀!!𝑠!!.

The ratio between these two rate constants is approximately 1.5, which does not

account for the seven-‐fold ratio of the overall rates of initiation. In addition, the first

RNAP binding rate constant on the nicked template matches the linear result,

2.1×10!𝑀!!𝑠!!, in the previous TIRF experiments (Friedman and Gelles 2012),

which is to be expected since they are both relaxed constructs. The rate of binding

the RNAP to the supercoiled versus nicked circular templates does not account for

the increase in overall rate of transcription initiation.

Closed Complexes

The next forward rate constant in the mechanism I wanted to determine was

the rate constant between the two closed complexes (𝑅𝑃!! ⟶ 𝑅𝑃!!). Since closed

complex formation does not require NtrC or ATP (Friedman and Gelles 2012), I

incubated the circular templates attached to the slide surface with 0.15nM of cy3-‐

𝜎!" RNAP alone (Figure 3.5A). The first experiment I performed to test the rate

constants of the closed complexes, was to measure the length of time the RNAP

stayed bound to the circular templates at 1-‐second per frame (Figure 3.5 B, C).

Since there are two closed complexes in the mechanism of transcription initiation,

34

the dwell times of the RNAP are expected to be a bi-‐exponential function (Friedman

and Gelles 2012). While the background signal interfered with the experimental

data, there was not much qualitative difference between the dwell times of the RNA

polymerases on supercoiled versus nicked templates.

Using the same experimental procedure as before (Figure 3.5A), the longer

dwell times were measured by increasing the time between frames to 5 seconds

(Figure 3.5 D, E). Unlike the 1-‐second per frame experiment, the background signal

in the 5-‐second per frame experiment did not interfere with the experimental data

from the nicked and supercoiled templates. Unfortunately, the bi-‐exponential fit did

not agree with the data and there was no reason that it should be fit to a tri-‐

exponential curve with three closed complexes. Qualitatively, however, the dwell

times on the supercoiled and nicked templates for the five-‐second per frame are

similar, agreeing with the data from the 1-‐second per frame experiment. Taken

together, the data from the 1-‐second and 5-‐second per frame experiments were not

strong enough to reach a conclusion on determining the rate constants between the

two closed complexes. To determine the rate constants involved in closed complex

formation, these experiments should be repeated with a different slide or re-‐labeled

circular templates to reduce the level of the background signal.

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Figure 3.5: Closed Complex Dwell Times. A. Diagram of experimental protocol. B. Cumulative frequency distribution of closed-‐complex lifetimes for the 1-‐second/frame experiment. C. Semi-‐logarithmic plot of data in B. D. Cumulative frequency distribution of closed-‐complex lifetimes for the 5-‐second/frame experiment. E. Semi-‐logarithmic plot of data in D. In B-‐E, the red circles correspond to the polymerases bound to the supercoiled templates (1-‐second per frame: n=241, 5-‐second per frame: n= 255), the blue circles correspond to the RNA polymerases bound on the nicked templates (1-‐second per frame: n=211, 5-‐second per frame: n=196), and the black circles correspond to the randomly chosen sites that did not contain a visible template (1-‐second per frame: n=850, 5-‐second per frame: n=942). The logarithms used here were loge (C-‐E).

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Open complex dissociation

In addition to measuring the rates of the forward reaction for transcription

initiation, I also wanted to determine the rate of open complex dissociation on the

supercoiled and nicked circular templates (𝑅𝑃! → 𝑅 + 𝑃). Open complexes were

formed in the same way as they were for the initial experiments on transcription

initiation (Figure 3.1A), except that instead of adding the NTPs after 45 minutes, the

lane was washed with buffer and oxygen scavengers (Figure 3.6A) to remove free

RNAP and allow the open complexes formed to dissociate. The positions of the open

complexes were viewed only once every 10 minutes to minimize photo bleaching.

At the first frame, I found all of the open complexes that were co-‐localized with

either the supercoiled templates or the nicked templates. The rate of open complex

dissociation on the supercoiled template was 1.3± 0.2 ×10!!𝑠!! and on the

nicked template was 1.6± 0.3 ×10!!𝑠!! (Figure 3.6B). Unlike the 𝜎!" results

(Revyakin, Ebright, and Strick 2004), these rates are not statistically different, which

indicates that there is no evidence that the 𝜎!" RNAP dissociates differently on the

supercoiled template compared to the nicked template.

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Figure 3.6: Dissociation of the Open Complex. A. Diagram of the procedure to determine the rate of open complex dissociation. B. Semi-‐logarithmic plot of the open complexes present as a function of time. The red circles represent the log of open complexes present on supercoiled templates (n=42) at a certain point in time, while the blue circles represent the log of open complexes present on nicked templates (n=43). The natural logarithm of the number of open complexes present was taken to create the graph. The solid lines represent single exponential fits on the data while the dotted lines describe the standard error on the fit (Equation 3.2).


TIRF Microscope and Slide Preparation

The TIRF microscope used in these experiments was the same as that

previously described (Friedman and Gelles 2012; Friedman, Chung, and Gelles

2006). The microscope slides were prepared with PEG and biotinated PEG on the

slide, and PEG on the coverslip (Friedman, Chung, and Gelles 2006). The slides were

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stored in a -‐80˚C freezer for up to two years after the PEG was added. They were

then cleaned with a Tris-‐Acetate (pH 8) buffer on the day of the experiment and four

lanes were created on the slide using grease. Both the supercoiled and nicked

circular templates were attached to the surface of the microscope slide using

streptavidin.

Solutions

All buffers containing a molecule attached to a fluorescent dye contained

oxygen scavengers (Friedman, Chung, and Gelles 2006). The buffer for the circular

templates, cy3-‐𝜎!" RNA polymerases, and probes were the same as previous

experiments (Friedman and Gelles 2012). The cy3-‐𝜎!" was stored separately from

the RNA polymerase core, both in a -‐80˚C freezer, and then combined in a 1.5:1 ratio

of Cy3-‐𝜎!": core, along with 0.28 mg/ml BSA, and 0.9 mM DTT just before adding to

the reaction mixture. Unless otherwise stated, the concentration of RNAP was

0.8nM, of NtrC was 14nM, of ATP was 2mM, of CTP and GTP was 0.5mM, and of the

cy5-‐probe was 10nM.

Fit Equations

The equation used to fit the cumulative plot of overall initiation and the time

until first RNAP binding (Figure 3.4 B, C) was

𝑓 𝑡 = (!"×!!!")×(!!!!!× !!!!"" !!"!!×(!!!")×(!!!!!×!!"")!

(3.1)

where Af is the fraction of the templates that were able to initiate (the active

fraction), n is the total number of templates present, Nz is the number of templates

with a co-‐localized probe at frame one, t is the time, R is the rate constant on either

the supercoiled or nicked template, and ROFF is the rate constant on the off DNA

39

locations (Friedman and Gelles 2012). The equation used to fit the single

exponential decay for the open complex dissociation (Figure 3.6B) was

𝑓 𝑡 = 𝑛×𝑒!! ! (3.2)

where n is the number of open complexes present at frame 1, t is the time, and R is

the rate of dissociation.

40


Rather than using linear DNA to measure the mechanism of transcription

initiation at the glnAp2 promoter, I created a circular template that could be used in

place of the linear DNA. The circular template was labeled with an Alexa Fluor 488

dye and biotin such that it could be seen by the microscope and attached to a

microscope slide. It also contained the necessary sequences for transcription

initiation and elongation for a 𝜎!" RNAP and had the ability to be negatively

supercoiled by the enzyme gyrase or relaxed by the enzyme Nt.AlwI. Using two

topoisomers of the circular template, the negatively supercoiled and the nicked

templates, I performed a series of single molecule TIRF experiments to determine

the mechanism for transcription initiation at the glnAp2 promoter. The rate

constants that I was able to determine are summarized in Figure 4.1. I found that

the overall rate of initiation on negatively supercoiled DNA is seven times faster

than on relaxed DNA, but not because of the first few forward rate constants in the

mechanism.

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Figure 4.1: Mechanism of Transcription Initiation for 𝝈𝟓𝟒-‐RNAP on Circular DNA. The rates in red are for RNA polymerases on supercoiled templates while the rates in blue are for RNA polymerases on nicked templates.

Throughout the TIRF experiments performed here to determine the

mechanism of transcription initiation, the background signal was higher than it was

for the previous experiments with linear DNA (Friedman and Gelles 2012). In the

case of the closed complex dwell times, the background signal interfered with the

results on supercoiled and nicked templates. I believe that the high level of

background signal was caused by the significant number of circular templates

attached to the surface of the slide that did not contain an Alexa Fluor 488 dye. The

amine-‐ester labeling reaction that I performed on the synthesized oligonucleotide

was only 20% efficient. Since I assumed that the cy5-‐probe could not be visualized

without elongation occurring on a DNA molecule, I determined the labeling

efficiency by finding the fraction of probes that co-‐localized with circular templates.

Another factor that may have contributed to the high background signal is that there

might have been nonspecific binding to the surface of the slide. Repeating the

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experiment to determine the dwell times of the closed complexes with a slide

prepared with an alternative method or using newly labeled oligonucleotide during

the creation of the circular templates may reduce the background signal.

While I was unable to determine all the rate constants in the mechanism of

transcription initiation for the circular templates, the preliminary data for the

𝑅𝑃!! ⟶ 𝑅𝑃!! step suggests that there is little difference between the rates on

negatively supercoiled and nicked templates. Therefore, the step that accounts for

the most difference between the rate constants of transcription initiation on

negatively supercoiled and nicked circular templates is the formation of the open

complex (𝑅𝑃!! → 𝑅𝑃!), which is also the rate limiting step in the reaction with

linear DNA (Friedman and Gelles 2012). If this is the case, then the rate of open

complex formation could be affected by negative supercoiling in two ways. First, the

negative supercoiling may reduce the force needed for the 𝜎!"-‐RNA polymerase to

open the DNA. The second is that the plectonemes formed in the negatively

supercoiled DNA may make it easier for the NtrC to interact with the 𝜎!"-‐RNA

polymerase thereby opening the double helix faster (Schulz, Langowski, and Rippe

2000). Finally it could also be a combination of both of these factors. To test how

supercoiling effects the rate limiting step in transcription initiation, I could use the

circular template described in chapter two, but rather than negatively supercoil it

with gyrase in the last step, I could positively supercoil the template with a reverse

gyrase. I could then compare the overall rate of transcription initiation on positively

supercoiled, negatively supercoiled, and nicked templates. Both the positively and

negatively supercoiled templates create plectonemes in the DNA, but unlike the

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negatively supercoiled template, the positively supercoiled template will make the

force to open the DNA greater rather than less than it would for the relaxed

template. Therefore, if the overall rate of transcription initiation on the positively

supercoiled template is the same as that on the negatively supercoiled template,

then the proximity of the NtrC to the 𝜎!"-‐RNAP accounts for the change in rate, not

the force needed to open the double helix. If however, the overall rate on the

positively supercoiled template is slower than it is on the nicked template, then the

reduced force needed to open the double helix explains the faster rate on the

negatively supercoiled template, not the accessibility of the 𝜎!"-‐RNAP to NtrC.

Finally, if the overall rate on the positively supercoiled templates is somewhere

between that on nicked and negatively supercoiled templates, then both the

proximity of the NtrC and the force needed to open the DNA account for the

increased speed of transcription initiation on the negatively supercoiled templates.

I found that the rate constants of 𝜎!"-‐RNAP open complex dissociation on

negatively supercoiled and nicked templates were the same within error. While

open complexes attached to negatively supercoiled DNA are expected to be more

thermodynamically stable, it apparently does not affect the dissociation rate of the

𝜎!"-‐RNAP. My finding contrasts with open complex dissociation of the 𝜎!"-‐RNAP on

supercoiled DNA, where the kinetic stability of the open complex does depend on

the superhelical density of the DNA (Revyakin, Ebright, and Strick 2004). However

in those results, the authors compared open complexes on positively supercoiled

DNA with those on negatively supercoiled DNA when performing their experiments,

rather than comparing the stability on negatively supercoiled and relaxed DNA that

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I use here. It would be interesting to determine if in my system the 𝜎!" RNAP has

similar open complex dissociation rates on relaxed and negatively supercoiled

templates like its 𝜎!" RNAP counterpart, or as the literature suggests, a differing

stability.

Using circular templates and single molecule TIRF microscopy, I have

described here the beginnings of the mechanism of transcription initiation using a

𝜎!" RNAP on negatively supercoiled DNA. More work needs to be done to

determine which is the step in the mechanism that accounts for the dramatic change

in overall rate between transcription initiation on the negatively supercoiled

templates compared to nicked templates, but preliminary data suggests that it is the

isomerization of the second closed complex to the open complex.

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