Evidence for Dna Oxidation in Single Molecule Fluorescence

EVIDENCE FOR DNA OXIDATION IN

SINGLE MOLECULE FLUORESCENCE STUDIES

A thesis presented to

the faculty of

the College of Arts and Sciences of Ohio University

In partial fulfillment

of the requirements for the degree

Master of Science

Douglas Wylie

August 2006

This thesis entitled

EVIDENCE FOR DNA OXIDATION IN

SINGLE MOLECULE FLUORESCENCE STUDIES

DOUGLAS WYLIE

has been approved for

the Department of Physics and Astronomy

and the College of Arts and Sciences by

Ido Braslavsky

Assistant Professor of Physics and Astronomy

Benjamin M. Ogles

Dean, College of Arts and Sciences

WYLIE, DOUGLAS, M.S., August 2006. Physics and Astronomy

EVIDENCE FOR DNA OXIDATION IN SINGLE MOLECULE FLUORESCENCE

STUDIES (103 pp.)

Director of Thesis: Ido Braslavsky

Single molecule fluorescence microscopy is a powerful tool to investigate local environments at the nanometer scale. Oxidation of DNA is an important problem that leads to DNA mutations. In this project, single molecule fluorescence measurements were used to study the interaction between covalently bound nucleotides (dNTP) and fluorophores (Cyanine dye), dNTP-Cy3 that have been incorporated into DNA. By tracing changes in the fluorescence signal, evidence for single events of DNA oxidation were found that could otherwise not be observed at the ensemble level. The differences between the nucleotides and their quenching properties are shown here especially guanine’s ability to temporarily quench the fluorescence of Cy3 completely. After some illumination time, the quenching of the dye by the guanine changes dramatically. We interpret this change to possible oxidation of the guanine base. This work could lead to a method for monitoring and investigating important DNA oxidation processes which are essential to Genomic and Cancer research.

Approved:

Ido Braslavsky

Assistant Professor of Physics and Astronomy

Table of Contents

Page

Abstract...... 3 List of Figures ...... 6 1 Introduction and Theory...... 8 1.1 Importance of understanding DNA damage...... 9 1.1.1 Single Nucleotide Polymorphisms (SNPs)...... 10 1.2 Causes of DNA damage and their effects ...... 12 1.2.1 Radicals ...... 12 1.2.2 Oxygen and its reactive derivatives, ROS...... 14 1.3 One-electron oxidation of DNA with emphasis on guanine ...... 16 1.3.1 Observation of site-selective Guanine-Oxidation...... 19 1.4 What is fluorescence? ...... 21 1.4.1 History of fluorescence ...... 21 1.4.2 Stokes’ shift phenomenon ...... 22 1.4.3 Jablonski energy diagram and quenching pathways ...... 24 1.4.4 Photobleaching and triplet-state reactions...... 26 1.5 Total Internal Reflection Fluorescent Microscopy (TIRFM)...... 31 1.6 Brief experimental outline and aim ...... 34 2 Sample Preparation and Experimental Method...... 35 2.1 Laser setup and TIR illumination ...... 35 2.2 Cleaning procedures and surface chemistry protocols ...... 39 2.2.1 Cleaning the glass slides...... 40 2.2.2 Application of polyelectrolyte multilayers...... 41 2.2.3 Surface activation for further attachments...... 42 2.2.4 Streptavidin application procedure ...... 43 2.2.5 Preparation and application of washing buffers...... 45 2.3 Properties of Cyanine Dyes, and DNA preparation...... 45 2.4 Observation of sample and data acquisition...... 51 2.5 Initial data analysis using custom developed correlation software ...... 54 3 Results and Comments ...... 56 3.1 Editing and analysis of movie data...... 56 3.2 Cy3-Primer/DNA by itself...... 58 3.3 dATP-Cy3/DNA...... 61 3.4 dCTP-Cy3/DNA...... 64 3.5 dUTP-Cy3-Primer/DNA (T404)...... 66 3.5.1 dUTP-Cy3/DNA (T407)...... 70 3.6 dGTP-Cy3/DNA...... 74 3.6.1 dGTP-Cy3/DNA (sample 2)...... 77 3.7 Preliminary results for incorporated dGTP-Cy5 with oxygen scavenger solution..80

Page

4 Discussion...... 83 4.1 Evidence for Guanine oxidation...... 83 4.2 New light on interpretation of photobleaching kinetics in single-molecule fluorescent traces...... 91 5 Conclusions and Suggestions for Future Work...... 92 6 References...... 95

List of Figures

Page

Figure 1) Ball and stick image of DNA double-helix...... 8 Figure 2) Chemical Structure of all 4 DNA bases along with Uracil present in RNA instead of Thymine...... 10 Figure 3) Diagram showing the π - stacking interactions among intrastrand bases and also hydrogen bonding between complementary base-pairs...... 14 Figure 4) An example of how oxidation of the original guanine base can induce mis-pairing of complementary bases...... 18 Figure 5) Structure of Cy3 (top) covalently attached to the de-oxy Cytosine Triphosphate nucleotide, dCTP (bottom)...... 20 Figure 6) Typical diagram showing Stokes’ shifted absorption and emission spectra...... 22 Figure 7) Diagram showing lowering of excited state singlet levels of a polar fluorophore due to solvent relaxation with surrounding medium...... 24 Figure 8) Jablonski diagram showing possible pathways for an excited electron and the various energy losses, transfer mechanisms and timescales for each transition ...... 25 Figure 9) Graph showing fluorescent intensity vs time for 3 single-molecules of Cy3 attached to a DNA-Primer sequence...... 27 Figure 10) Simplified Jablonski diagram showing various reaction rates and photobleaching behavior ...... 29 Figure 11) Picture showing path taken by laser light through objective and reflected at cover glass/sample surface interface ...... 32 Figure 12) Nice diagram depicting approximate penetration depth d, of the evanescent wave at an incident angle of 62 degrees for a 542nm wavelength laser beam…...... 33 Figure 13) Microscope with red (top left) and green (middle left) laser set-up on air-cushioned table ...... 35 Figure 14 a and b) Absorbance and emission spectral profiles for the cyanine dyes, Cy3 and Cy5, along with their appropriate filter cubes allowing maximum desired light through with minimal overlap of unwanted light ...... 36 Figure 15) Plot showing relative intensity of evanescent wave against displacement from surface, z (nm) and angle of incidence, θ (degrees) for 532nm wavelength excitation ...... 39 Figure 16) Diagram showing surface preparation prior to DNA attachment ...... 44 Figure 17) Chemical structure of Cy3-dCTP and Cy5-dUTP...... 46 Figure 18) DNA templates, named T404 and T407 both with identical primers (in bold) shown underneath as complementary sequences growing from the 3’-end of the template, Oligos provided by Helicos Biosciences...... 47

Page

Figure 19) DNA with incorporated nucleotides attached to glass surface...... 50 Figure 20) Separated Laser and filter module with 2 pinhole apertures...... 53 Figure 21) Showing amount of correlation from 1st movie frame using the red laser with all subsequent frames using the green laser...... 55 Figure 22) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of Cy3-DNA/Primer...... 58 Figure 23) Selected frames from Cy3-Primer/DNA movie illustrating bleaching behavior over time ...... 60 Figure 24) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dATP-Cy3-DNA ...... 61 Figure 25) Selected frames from dATP-Cy3/DNA movie showing decrease in Fluorescent S-Ms due to Photobleaching...... 62 Figure 26) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dCTP-Cy3-DNA ...... 64 Figure 27) Selected frames from dCTPCy3/DNA movie showing decrease in fluorescent S-Ms due to photobleaching...... 65 Figure 28) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dUTP-Cy3-DNA ...... 66 Figure 29) Selected frames from dUtpCy3/DNA movie showing decrease in fluorescent S-Ms due to photobleaching...... 68 Figure 30) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dUTP-Cy3/DNA (T407)...... 70 Figure 31) Selected frames from dUTP-Cy3/DNA (T407) movie showing decrease in fluorescent S-Ms due to photobleaching...... 72 Figure 32) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dGTP-Cy3/DNA...... 74 Figure 33) Selected frames from dGTP-Cy3/DNA movie showing decrease in fluorescent S-Ms due to photobleaching...... 76 Figure 34) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dGTP-Cy3/DNA (sample 2) ...... 77 Figure 35) Selected frames from dGtpCy3/DNA_sept movie showing decrease in fluorescent S-Ms due to photobleaching...... 78 Figure 36) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dGTP-Cy5/DNA in O2 scavenger...... 80 Figure 37) Selected frames from dGtpCy5/DNA_O2_scavenger movie showing decrease in fluorescent S-Ms due to photobleaching...... 82 Figure 38) Diagram showing 11 of the main guanine adducts and oxidative lesions formed via one-electron oxidation, hydroxyl radical damage and 1O2 mediated reactions in solution...... 84 Figure 39) Diagram depicting possible reaction pathways explaining observed S-M fluorescence profiles of Cy3-dGTP...... 86 Figure 40) Diagram showing in more detail the electron transfer process between an acceptor and donor molecule...... 87

1 Introduction and Theory

Although this thesis is within the department of physics program the main body of research is in single molecule fluorescence studies of various DNA-dye configurations. It is therefore necessary to give a slightly more bio-chemically orientated introduction to explain the significance of DNA itself and why it is so important to understand as much about it as possible. Below is a typical image of a segment of DNA.

Figure 1) Ball and stick image of the DNA double-helix

Note the twisted ‘ladder’ structure with sugar-phosphate backbones (red and blue) running anti-parallel to each other and the horizontal ‘rungs’ known as complementary base pairs, Adenine (purple) with Thymine (green) and Guanine (yellow) with Cytosine (cyan). Figure taken with permission from http://www.chemicalgraphics.com/paul/DNA.html

1.1 Importance of understanding DNA damage.

DNA is probably the most essential and one of the most studied molecules in the entire human body. With the completion of the Human Genome Project (HGP) [1] and the technological advances made in imaging techniques especially in the area of single- molecule (SM) fluorescence microscopy,[2-7] the ability to probe ever further and in more intricate detail the inner workings and interactions of this amazingly complex molecule is being realised. DNA interactions with other molecules in the cell environment and how the sequence of bases make up the various genes which code for the amino-acids and therefore proteins, is being better understood at the molecular and even sub-molecular level. It is interesting to note that less than 5% of the 3 billion base sequence, i.e. the order of the four nucleotides ATCG in DNA, is actually used in coding for proteins [8]. The other non-coding regions are thought to ensure structural integrity and control the amount of protein formed. Recently it has been suggested that certain ‘junk’ regions of DNA might in fact serve as potential sacrificial traps for damage causing radicals and oxidizing electron ‘holes’ that could otherwise migrate to the more important gene-coding regions of DNA [9]. There is still therefore much to learn and now we have the tools and the foundation, from the HGP, to undertake the necessary experiments to answer more of these questions.

Figure 2) Chemical Structure of all 4 DNA bases along with Uracil present in RNA instead of Thymine.

Diagram taken from http://homepages.ius.edu/GKIRCHNE/bases.jpg

It is well known that damage caused to DNA such as mutations (defined as a variation in the DNA sequence occurring in <1% of the population), single nucleotide polymorphisms (SNPs), strand breaks and cross-linking between DNA and protein can lead to ageing, cell death, cancer and other diseases [10-16]. To understand what causes these defects, how they occur and what damage they actually do to DNA is therefore very important.

1.1.1 Single Nucleotide Polymorphisms (SNPs)

A SNP is merely a single nucleotide base variation in the DNA sequence. In other words a change in the sequence ATTCCGGCA to ATTCTGGCA is considered a SNP if it occurs in >1% of the population SNPs occur on average every few hundred bases and as such there can be as many as 10,000,000 SNPs [8]. They are effective markers along the DNA strand which prove to be very helpful in finding genes thought to be involved in various diseases. Up to 90% of all human genetic variation arises from SNPs and, as with

11 mutations, most have no effect but some are believed to increase susceptibility to disease and also alter the efficiency of the cell’s response to drugs. The harmless or un- interesting SNPs occur mostly in the non-coding regions of DNA and therefore are not closely associated with any genes related to a particular disease. However there are of course some SNPs that do occur within the gene coding region of a section of DNA and while most of them do not cause the disease themselves some are thought to alter slightly the specific function of an associated protein. This is because the original DNA sequence of interest has been changed by one base which means that one amino acid could incorrectly be coded for thereby altering slightly the overall structure of the related protein.

An example of how SNPs are an important marker for improving potential disease detection and effectiveness of the prescribed medicine is shown in the case of Alzheimer’s. One of the genes linked with alzheimer’s is called apolipoprotein E or Apo E. It contains two SNPs which therefore give rise to three alleles (alternate forms of a gene) of the gene, E2, E3, and E4. The three alleles only differ by one base each and also by one amino acid each. It has been shown that those who inherit one or more of the E4 alleles have a greater chance of getting Alzheimer’s compared to those inheriting the E2 form. It therefore seems that the change of this particular amino acid in the protein increases the likelihood of developing Alzheimer’s. One should note of course that this is not a 100% hard and fast rule. A person with both E4 alleles might never get alzheimer’s just as a person with two E2 alleles can equally develop the disease. This is because Apo E is just one of several known genes associated with alzheimer’s. It is these multi-gene variations common among serious conditions such as heart disease, diabetes and cancer that make genetic testing to find them extremely difficult [1, 8]. This is why a full and easily obtainable SNP map along with large population studies and cross checking among them will be of great benefit in developing more personalized and effective medical diagnoses and treatment. This leads us to ask the question, what are the possible causes of these types of variations in the DNA and the damage that they can inflict?

1.2 Causes of DNA damage and their effects

1.2.1 Radicals

Major sources of DNA damage come from the formation of radicals. A radical (sometimes called a "free radical") is a molecule that has an unpaired electron (represented by a dot next to the chemical structure, e. g. A.). Some radicals are stable

and long-lived (the most common example is O2, the oxygen in the air we breathe, which is a "di-radical" as it has two unpaired electrons), but most radicals are highly reactive and are usually very unstable.

Due to its very nature a radical needs to pair its unpaired electron with another electron, and will react with another molecule in order to obtain this missing electron. If a radical achieves this by ‘stealing’ an electron from another molecule then that other molecule also becomes a radical (Reaction 1), and thus a self-propagating chain reaction begins (Reaction 2).

If a radical pairs its unpaired electron by reacting with a second radical, then the chain reaction is terminated, and both radicals ‘neutralize’ each other (Reaction 3).

Radicals are produced by normal aerobic (oxygen-requiring) metabolism and are necessary to life. They are used by the immune system to fight off harmful bacteria and viruses. However, excess amounts of radicals are harmful because of their high reactivity. They can be formed easily by compounds that readily give up a single electron (e.g., nucleotide bases, principally guanine, and polyunsaturated fatty acids). Radicals are also

13 produced by processes other than normal metabolism such as from UV and ionizing radiation, smoking and other pollutants, herbicides and pesticides [13], and can even be found in certain types of food (e. g., deep-fat fried foods). Radicals can also damage lipids (the molecules which make up cell membranes) and proteins as well as DNA. It is a consequence of these extra sources of radicals being introduced to the human body along with endogenous sources that can overwhelm the natural repair mechanisms and enzymes especially under periods of oxidative stress. This then results in a net build up of harmful reactive radicals that are thought to increase the rate of cell death, aging and susceptibility to chronic diseases such as cancer. Radical damage is significant because it can proceed, as mentioned previously, like a chain reaction due to the unpaired electron ‘desperately seeking’ and often stealing a single electron from another nearby molecule. This ‘stealing’ of single electrons modifies the base and sugar molecules within a DNA strand and thereby alters their structural and electro-chemical properties such as the ability of a nucleotide to bind effectively with its complementary base i.e. adenine with thymine, A-T, and guanine with cytosine, G-C. This type of normal healthy base association is otherwise known as Watson-Crick pairing [17].

Transversion is the process where a normal b-p for example G-C mutates to a T-A or C-G pair depending on the type of radical species formed during the reaction [18]. This will be explained in more detail in the next section. After a base has been modified by a radical it too can become a radical either having gained or lost an un-paired electron. Now this base radical can undergo a further reaction and become irreversibly oxidised or damaged leading to the mismatch of complementary bases or even no complementary base at all [19-21]. This increases the probability of permanent damage whether it is a break in the DNA strand (usually at the particular sugar molecule involved [20, 22, 23] or a change in the structural integrity and electron transport properties of the DNA (predominantly in the π - stack [24],[25, 26] and also in the hydrogen bonds between opposite bases [18, 27] because of the mutation in the original sequence at that site.

Figure 3) Diagram showing the π - stacking interactions among intrastrand bases and also hydrogen bonding between complementary base-pairs.

Figure taken from reference [18] K. Kawai, T. Majima / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 3 (2002) 53–66

A key component in the formation of radicals among DNA bases and the sugar- phosphate backbone is oxygen or more specifically, Reactive Oxygen Species known as ROS.

1.2.2 Oxygen and its reactive derivatives, ROS.

The oxygen in the air we breathe is, a free radical or more precisely a di-radical, as it has two unpaired electrons [28, 29]. It is normally found in its ‘ground’ (not energetically 3 excited) state and is symbolized by the abbreviation O2. Molecules whose outermost pair of electrons have parallel spins (denoted by ) are in what’s known as the ‘triplet’ state and those molecules whose outermost pair of electrons have anti-parallel spins (denoted by ) are in the ‘singlet’ state. Spin can either be up, +1/2 or down, -1/2 and for an

15 electron pair with anti-parallel spin , i.e. S= + ½ - ½ =0, there are 2S+1 states giving rise to the multiplicity (number of possible orientations of spin angular momentum) value of 1, hence the term singlet state. Similarly the unpaired electrons with parallel spin will have total spin, S=1 and a multiplicity value of (2x1) + 1=3, hence triplet state. 3 Ground state oxygen is in this triplet state (indicated by the superscripted "3" in O2). Its two unpaired electrons have parallel spins which means that, according to the Wigner spin conservation rule, these electrons are not allowed to react with most other molecules in their singlet states due to ‘spin forbidden’ or highly improbable transitions. Thus, ground-state or triplet oxygen is not very reactive. However, if triplet oxygen absorbs sufficient energy (e.g. by photosensitization) [30-32] to reverse the spin of one of its unpaired electrons, it forms the excited singlet state. This excited singlet oxygen molecule is one of several very reactive oxygen moieties including radicals and are collectively known as reactive oxygen species, (ROS).

1 Singlet oxygen, abbreviated O2*, has a pair of electrons with opposite spins and is therefore not a radical although it is highly reactive. The * symbol is used to indicate that this is an excited state with excess energy. An important point to remember is that neither triplet nor singlet state molecules are necessarily in the excited state. The designation of ‘triplet’ or ‘singlet’ refers only to the spin state.

This reaction can also be written in this form:

There are other types of ROS such as the extremely reactive hydroxyl radical, OH·,

and the less reactive and sometimes necessary superoxide radical, O2 among others.

These have important consequences in biological systems where other essential intermediates such as transition metals like Fe (frequently found in cells) and Cu facilitate their production within the cell where they can do most damage. However, with regards to our experimental set-up it is thought that the main ROS produced is the reactive singlet oxygen. This is because molecular ground state oxygen only requires about 0.98eV of energy to be excited to the more reactive singlet state and this can also come from the green laser. Another source of singlet, ¹O2, oxygen is from possible triplet state reactions between the Cy3 dye and the oxygen within the buffer solution [30]. This has the added effect of returning the triplet dye to the singlet ground state ready for re- excitation again. Also, the lack of transition metal ions such as Fe2+ present in the surrounding buffer solution would greatly diminish the amount of OH· formed.

1.3 One-Electron Oxidation of DNA with emphasis on Guanine

DNA oxidation is a widely studied and very important area of research into understanding the mechanisms behind what causes cancer and ageing [33]. There are over 100 known oxidative by-products, or adducts, that can be formed within the cell and DNA as part of reactions with radicals and other ROS [34]. In any one day a normal body can absorb 10^12 O2 molecules and it’s thought that 1-2 out of every 100 of those O2 molecules end up being involved in some form of oxidative radical inducing reactions that can damage the nucleotides and the sugar-phosphate backbone [20, 22] as well as lipid membranes and other organelles within the cell. In the case of DNA all four bases can be oxidised (lose an electron) quite easily because of their relatively low oxidation potentials [35] . Even though the redox potentials of the nucleotides change slightly from being free nucleotides in aqueous solution, [35] to the formation of single stranded and double stranded DNA, the relative order of oxidation is still similar with Guanine being the easiest to oxidise followed by Adenine, Cytosine, Thymine and finally Uracil. The

17 oxidation potentials of the 4 bases and uracil in solution relative to the normal hydrogen electrode (NHE) are dG:1.47V < dA:1.94V < dC:2.12V and dT:2.09V < U:2.37V [36] . Using molecular orbital theory calculations Colsen et Al showed that base-pairing with Cytosine reduced the overall ionization potential of the guanine by approx. 0.54eV [37, 38]. They also proposed from their calculations that the hydrogen bonds are strengthened within a G-C ion pair compared to the neutral parent G-C base pair. Also, Majima and Kawai more recently [18] have shown experimentally that the oxidation potential of guanine decreases by approximately 100mV upon base-pair binding with cytosine thus facilitating guanine’s oxidation. They also attributed this to the hydrogen bonding between the bases. Interestingly there is also an associated proton transfer [39] from the guanine to the cytosine coupled with the one-electron oxidation. This would have the effect of neutralizing the charge on the guanine radical cation having now given up a proton along with the oxidised electron. In fact, this guanine moiety is known as a neutral guanine radical. Guanine can also be oxidised to several other forms [31, 40] with the most common (and probably most studied) being the guanine adduct 8’oxo-dG or 8oxoG which itself is easily oxidised and is associated with the transversion of G-C to A-T mismatching as mentioned earlier.

The reason why guanine oxidation is of such interest comes from the fact that, even though all of the other bases can be oxidised to some degree or another, the radicals formed tend to ‘migrate’ to nearby guanine bases or double (5’-GG-3’) and even triple (5’-GGG-3’) guanine sites along the DNA chain [13, 41] . This migration of charge from the radical travels along the π-stack [12, 24, 26, 42-50] (de-localised pi-molecular orbitals along the base sequence) on an intrastrand pathway until it ‘falls’ into a ‘trap’ at the particular guanine or multiple guanine site. Again this is because of the lower oxidation potential of guanine compared to the other bases so the negative electron donated from the guanine neutralizes the migrating radical but in so doing causes itself to become a radical and prone to further damage.

The oxidized forms of guanine are known as lesions and these lesions are one of the main causes in the breakdown in integrity of the DNA replication process and increased mutation rates.

Figure 4) An example of how oxidation of the original guanine base can induce mis- pairing of complementary bases

Figure again taken from reference [18] K. Kawai, T. Majima / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 3 (2002) 53–66

1.3.1 Observation of site-selective Guanine-Oxidation

Being able to observe, directly or indirectly via a closely attached reporter molecule (such as a fluorescent dye), the oxidation of a specific single guanine base greatly increases our ability to understand the mechanisms behind these processes that do damage to the bases. Any new possible methods for observing specific guanine oxidation especially at the single-molecule level is therefore of valuable interest and well worth pursuing. Due to advances in the attachment of fluorescent molecules to individual nucleotides it has become possible to covalently bond the two together.

In previous studies many fluorophores have been attached via a linker arm of some few atoms in length to the respective nucleotide. Even though only a few atoms separated the two entities the ability to detect any change in electronic structure of the desired base would be difficult as only the outer electrons of the base are involved in any radical forming reactions. Therefore the linker arm itself could act as a shield and reduce the interaction of the fluorophore with the base. It must be noted that in a lot of cases depending on the experiment and variables to be observed it is actually desirable to have this shielding effect. Very strong quenching, especially from guanine [51-54] can drastically reduce the fluorescence of a particular fluorophore making it harder to detect at low concentrations. Care needs to be taken in comparing fluorescent profiles of various dye-nucleotide combinations so as not to attribute variations in intensity to an ambiguous factor. This is why a lot of research has been done on the various fluorescent and electro- chemical properties of dyes to enable experimenters to choose the most appropriate dye with the most suitable spectral profile. For example, in the field of single-molecule DNA sequencing [55-59] it is necessary to have reliable fluorescence data that is well understood so that the differences in profiles between the bases and their attached fluorophores as they are sequenced do not get misinterpreted. Next I will discuss nucleotides labelled with Cyanine -3 dye. [Figure 5] The cyanine dyes are known for being bright fluorophores and are used in many DNA applications.

Figure 5) Structure of Cy3 (top) covalently attached to the de-oxy Cytosine Triphosphate nucleotide, dCTP (bottom).

Above figure taken from http://las.perkinelmer.com/content/TechnicalInfo/nel576.pdf

The fact that the fluorophore (Cy3) is covalently bonded to the individual free nucleotide means that it has a much closer interaction with it due to the sharing of the outer electrons (i.e., those involved in the one-electron oxidation process). Any effects from the transfer of electrons, formation of radicals or oxidative reactions from the surrounding medium will have a much greater influence on the fluorescent profile of the attached Cy3 dye.

Now it is time to turn our attention towards the topic of fluorescence itself and how we can use this phenomenon to observe dye-nucleotide interactions at the single- molecule level.

1.4 What is fluorescence?

Fluorescence belongs to a wider group of light related phenomenon known as luminescence. Luminescence is the general term for any light that is emitted from a “cool” source and not from a hot body or other form of incandescent light. It is caused by the transition of electrons in an excited or higher energy state within a molecule to a lower less energetic or ground state. There are several other types of luminescence such as chemiluminescence (obtained from certain chemical reactions), bioluminescence (such as that given off from glow-worms and fireflies) and phosphorescence (the ability of a substance to “glow” even after the stimulated excitation has stopped, e.g. those glow in the dark stars on your bedroom ceiling as a kid!). If the source of luminescence comes from some form of electromagnetic radiation such as ultraviolet light or even other sources like pressure or mechanical deformation and it immediately (or within an extremely short time period i.e. a few nanoseconds) ceases to luminesce once the excitation source has stopped then this is known as fluorescence [60]. Also, in fluorescent materials the excited state has the same spin as the ground state. In other words transitions only occur from singlet (excited) states to singlet (ground) states and singlet-triplet interactions are rare or spin forbidden as previously mentioned.

1.4.1 History of fluorescence

As far back as 1500 BC the Chinese are reported to have written books on the observance of fluorescence and phosphorescence but it is more widely accepted, at least in the western world, that the discoverer of fluorescence was a German Jesuit priest named Athanasius Kircher born in 1602 in Giesa a.d. Ulster, Germany. This all round philosopher and scientist stumbled upon an interesting effect by injecting water on the wood extract lignum nephriticum and shining light on it. He was able to observe blue

22 reflected light and yellow transmitted light. It is this reflected blue light that is in fact fluorescence. One of his books entitled “Ars Magna Lucis et Umbrae” was published in 1646 in Rome and covers this plus many other observations and theories on light and, amongst other things, the motion of the planets!

1.4.2 Stokes’ shift phenomenon

It wasn’t until 1852 when Sir George Stokes first coined the term fluorescence which

was believed to have come from fluorite (Ca F2), a strongly fluorescent mineral. He was the first to observe that the excitation wavelength used was shorter than the emitted wavelength of light. This phenomenon was named after him and is known as Stokes’ shift.

Figure 6) Typical diagram showing Stokes’ shifted absorption and emission spectra.

Diagram taken from http://en.wikipedia.org/wiki/Stokes_shift

The reason for the shift to longer wavelengths comes from the decrease in energy of the excited molecule as it almost immediately (<10−¹²s) loses energy by vibration. This is known as vibrational relaxation. Once at the lowest vibrational state of the excited state level S1 a now lower energy photon is released enabling the electron to return back to its ground state, S0. After returning to its ground state the electron can then be excited again to a higher level and the process is repeated. In theory this cycle can go on indefinitely but in practice there are other factors that influence the particular energy pathway that the electron can take. There are also certain interactions that occur between the fluorophore and the coupled, or ‘tagged’, nucleotide that can greatly alter the fluorescence profile. In the case of Cy3 and Cy5, which are polar molecules [61-65], solvent effects play a significant role in the stokes’ shift of the dye. This is because an electron only takes on the order of a few femto-seconds to jump to a higher electronic state thereby altering slightly the molecular structure of the outer shells and changing its dipole moment. This process happens so fast that it takes time for the surrounding polar solvent water molecules to rearrange themselves and hence energy is transferred as the dye relaxes to the lowest exited state level. This solvent relaxation effect can occur over slightly longer time periods, approx. 10−10 secs which is significantly slower than the few pico-seconds it takes for normal vibrational relaxation. The energy level from which the photon is emitted will be reduced further and is closer to the ground state energy level leading to a larger stokes’ shift.

Another important consequence from this lowering of the excited state energy level is that it is also closer to the triplet state energy level and could therefore increase the rate of intersystem crossing between the spin states of the molecule. Because oxygen is naturally in its triplet state there is a greater chance of triplet-triplet dye-oxygen interactions. The triplet-triplet interaction converts the oxygen to singlet oxygen while the dye returns to the singlet ground state simultaneously.

In turn the reactive singlet oxygen is able to interact with both the dye itself (in terms of photobleaching) and also the nearby nucleotide causing possible forms of oxidative damage. A diagram showing the lowering of the excited state level due to solvent relaxation is shown below.

Figure 7) Diagram showing lowering of excited state singlet levels of a polar fluorophore due to solvent relaxation with surrounding medium.

Figure taken from http://micro.magnet.fsu.edu/primer/java/jablonski/solventeffects/index.html

1.4.3 Jablonski Energy Diagram and quenching pathways

The other possible pathways for the energy of an excited electron to take are best shown in what is known as a Jablonski diagram [66]. See figure 8 on following page.

Figure 8) Jablonski diagram showing possible pathways for an excited electron and the various energy losses, transfer mechanisms and timescales for each transition.

This figure is taken from http://micro.magnet.fsu.edu/optics/timeline/people/jablonski.html

The vast majority of molecules at room temperature tend to be in their ground state normally and any typical thermal kicks they might get from their surroundings isn’t usually enough to excite them to a higher energy state but merely ‘nudges’ them a few vibrational levels higher at most within the ground state. This is one of the main reasons why fluorescence can be such a powerful imaging technique in the first place. It should also be noted that most electrons under excitation within the visible wavelength spectrum only have enough energy to jump to the 1st excited state which means we can neglect any

S2 transitions in our experiment.

As can be seen from figure 8) once at the lowest level of the S1 state the electron can either go one of several ways depending on its surroundings and the various transition probabilities pertained to it. It can, after a few nano-seconds, spontaneously emit a photon and decay to a vibrational level in the ground state, this is fluorescence. In terms of singlet state to singlet state transitions, it may also decay back down to the ground state through various vibrational levels by internal conversion otherwise shown as non- radiative processes in the above Jablonski diagram.

The excited electron can also be ‘quenched’ by transferring its energy to another molecule as in ‘Fluorescent Resonant Energy Transfer,’ FRET [64, 67-69] or by colliding with molecules in the surrounding medium. It can also be quenched via electron transfer to an acceptor molecule in a process known as photoinduced electron transfer [70-72]. In some cases the quenching can be so pronounced that it stops the fluorescence completely as will be shown later in the case of Cy3 attached to the guanine nucleotide, dGtpCy3. In fact it is this complete quenching of fluorescence that leads to evidence for guanine oxidation. Once the guanine has been irreversibly oxidized or damaged, the fluorescence of Cy3 then starts and keeps fluorescing until it is eventually photobleached as well.

1.4.4 Photobleaching and triplet-state reactions

Another important transfer pathway is what’s known as intersystem crossing between the singlet spin state and its triplet state. As mentioned before in section (1.22) triplet state reactions are rare compared to singlet-singlet reactions but over time they do build up due to their much longer excited state lifetime (micro-milliseconds). This enables the triplet-state of the dye more time to undergo possible chemical reactions with surrounding molecules, most notably molecular oxygen, leading to photo-induced

27 chemical damage and changes in its covalent molecular structure [73]. When these changes cause the fluorophore to cease fluorescing it is said to have been photobleached. This is an irreversible process and in ensemble experiments the photobleaching of the dye causes an exponentially related decrease in the fluorescent intensity of the illuminated sample. However, at the single-molecule level this is characterized by a sudden cut-off in the fluorescence intensity profile [6, 65, 74-76]. A typical fluorescent intensity profile of a single bleached fluorophore, Cy3, is shown below.

Figure 9) Graph showing fluorescent intensity vs time for 3 single-molecules of Cy3 attached to a DNA-Primer sequence.

Single-Molecule Traces Cy3P4T4

200 S-M# 1 150 S-M#26 S-M#79 100 50 Intensity (a.u) 0 0 50 100 150 200 250 Time (seconds)

Figure taken from actual experimental results. One can clearly see the sudden cut-off of fluorescence which is a signature of irreversible photobleaching of the Cy3 fluorophore.

The exponential decrease in fluorescence can be either mono- or multi-exponential depending on the base-dye combination used [75] and is described by the following equation:

− t a τ F()tF= i ei (0) ∑τ eqn.[1] i i where, F(t) is the total fluorescent intensity at time t, F(0) is the intensity at time t=0, ai is a weighted factor accounting for the fluorescent quantum yield in a sub-population i of τ excited fluorophores, and i is the photobleaching lifetime for sub-species i. For a bi- exponential fit, as will be shown in the case of dGTP-Cy3, the variables a1 and a2 add up τ to 1. We can then group F(0) , ai and i together into two separate constants for each exponential and the equation then becomes:

−−tt ττ =+12 F()tFeFe12 eqn.[2] a a where, F = F 1 and F = F 2 1 (0) τ 2 (0) τ 1 2 The above equation can be understood in more detail by applying Michelson-Morley type reaction kinetics to the various molecular states and their transition rates [74]. Again this can be shown in a Jablonski diagram and by applying the appropriate reaction kinetics.

Figure 10) Simplified Jablonski diagram showing various reaction rates and photobleaching behavior.

There is an accumulation of photobleached product P formed from triplet state reactions. Here, kic is the

internal conversion rate, kEX is the excitation rate from the ground state to the excited singlet state, kF is the fluorescence emission rate, kisc is the intersystem crossing rate, kT is the triplet-ground singlet

transition rate and kb is the bleaching rate. Figure modified and taken from ref [74] E. Fureder-Kitzmuller et al. / Chemical Physics Letters 404 (2005) 13–18

The population dynamics in each of the four states can be described by the following rate reactions:

•

1) S0 = − kEX S1 + kT T + kF S1 + kic S1

•

2) S1 = kEX S0 − kF S1 − kic S1 + kisc S1

•

3) T = kisc S1 − kT T − kb T

•

4) P = kb T

where the steady state population condition is such that, S0 + S1 + T + P = 1

The excitation rate, kEX is proportional to the intensity of the illumination at the sample ε −1 surface, I (W/cm²), the molar extinction co-efficient, m (Mole¯¹ cm ), which is related to the absorbance area of the dye at a certain wavelength and inversely proportional to the photon energy hc / λ at the excitation wavelength. For Cy3 the extinction co-efficient is 150000M¯¹ cm−1 and the fluorescent photon has an energy of app. 2.25eV. One can solve the above rate equations by assuming that the molecular system, S0 , S1 and T are in

• • • equilibrium with each other therefore, S0 , S1 and T are all equal to 0.

By definition the relative amount of bleached photoproduct, P, at time t=0 has to be 0 and conversely as t →∞all of the dye will have been bleached. This leads to the simple relation:

− t τ P = 1 − e bl eqn.[3] τ τ where, bl is the bleaching lifetime as in eqns. 1 and 2, bl is given by the following rather lengthy reaction rate:

kkk++ ()()kkk++ kk + 1 τ isc T b + FiciscTb bl =, see [74-76] for a more in depth kkbisc kk bisc k EX and rigorous explanation.

As will be discussed in the results section, for all of the dNTP-Cy3 configurations, apart from dGTP-Cy3, a mono-exponential fit was sufficient to describe the bleaching rate as there were no obvious significant differences compared to that of a bi-exponential fitted curve. On the other hand the guanine/dye analog when incorporated into the DNA primer was by far the most quenched of the four nucleotides and exhibits a very different bleaching pattern to the others.

Factors influencing the rate of photobleaching include; the intensity of illumination used i.e. the higher the intensity the faster the bleaching rate, the type of fluorophore used with regards to its resistance to bleaching and the amount of molecular oxygen within diffusion distances of the fluorophore. Reactions involving light and oxygen that lead to photobleaching are labeled as photodynamic and the ROS given off from the triplet- triplet dye-oxygen interactions can damage nearby molecules such as nucleotides and especially guanine as is proposed to be the case in our experiment.

Due to the low illumination intensities required in order to obtain longer periods of fluorescence before bleaching it is necessary to have a highly sensitive camera and appropriate fluorescent microscopy set-up. Also, because of the long illumination periods under the laser the sample must be fixed in position which therefore requires it to be attached close to the surface of the coverslip. With these constraints in mind the most suitable type of sample illumination to use is ‘Total Internal Reflection’ or TIR. This is utilized in a type of microscopy technique known as ‘Total Internal Reflection Fluorescent Microscopy’ (TIRFM) [73]. It is the method used in this experiment and will be explained in more detail in the next section.

1.5 Total Internal Reflection Fluorescent Microscopy, TIRFM

There are many other types of optical imaging techniques such as confocal microscopy, fluorescent correlation spectroscopy (FCS), epi- and wide-field illumination microscopy and each have their own advantages and disadvantages depending on the type of illumination technique required and the particular variables under investigation. For this experiment TIRFM is most suitable because of the low excitation volume illuminated in the sample which greatly reduces any excess background fluorescence [77-80] from any freely diffusing fluorophores not attached to the surface. This reduction in

32 background noise is essential for efficient single-molecule detection. The low illumination depth, is possible due to the fact that above a certain critical angle of incidence between 2 mediums of different indexes of refraction, n1 and n2, an incoming light beam will be totally internally reflected at the boundary between the higher indexed medium, e.g. glass coverslip, n2=1.52 and the lower indexed medium e.g. sample specimen, n1=1.34.

Figure 11) Picture showing path taken by laser light through objective and reflected at cover glass/sample surface interface

Picture taken from http://www.microscopyu.com/articles/fluorescence/tirf/tirfintro.html

All the energy of the incident wave is reflected at the boundary. A standing wave is set up at the interface which then penetrates very slightly into the lower indexed medium. There is an electromagnetic field which propagates parallel to the incident plane and this is called an evanescent wave [5, 79]. It decays exponentially away from the surface and normal to the surface with a typical length of 150 nm, see equation 5.

The intensity of the evanescent wave is given by the equation: = − z / d I z Ie0 eqn [4] where, z is the distance normal to the surface into the lower indexed medium, Io is the intensity at the surface boundary and d is the characteristic penetration depth given by:

λ − dn=−0 [sin22θ nn 21/2 ] eqn [5] 4π 122c λ θθ> θ where, 0 is the light wavelength in a vacuum, c , c is critical angle of 61 degrees for glass/buffer solution boundary and n1 , n2 are refractive indexes of the glass slide and solution respectively. The depth, d is independent of the incident light polarization. For a green wavelength, 543 nm incident at an angle of about 62 degrees the penetration depth is approximately 200nm. This can be seen in the figure below.

Figure 12) Nice diagram depicting approximate penetration depth, d, of the evanescent wave at an incident angle of 62 degrees for a 542nm wavelength laser beam.

Figure taken from http://micro.magnet.fsu.edu/primer/java/tirf/penetration/index.html

Only those fluorophores close to the surface within the excitation volume are excited and this leads to higher signal to background ratio compared with epi- and wide-field illumination techniques with possible S/N ratios > 10 [80]. TIRFM is thus a highly sensitive and practical technique for imaging molecules close to or fixed to a surface such as streptavidin bound DNA strands [57] and is also well suited for studying biological phenomena at cell membrane boundaries, another important area of biological research.

1.6 Brief experimental outline and aim

Few experimental steps are outlined here. First single stranded DNA is annealed to primers labeled with Cy5 at their 5’- ends. Then single Cy3 covalently bonded nucleotides are incorporated onto the DNA primed template. After diluting to 25pM the DNA/nucleotide sample is washed into a flow cell and attached to the chemically treated surface via a streptavidin biotin linkage. After observing all four DNA/nucleotide configurations at the single-molecule level using TIRFM, I will show that the nucleotide de-oxyguanosine triphosphate, dGTP, completely quenches Cy3 fluorescence for various lengths of time until at some point later, fluorescence is suddenly initiated. Eventually photobleaching of the dye occurs upon continued illumination.

The consequences of this severe quenching by dGTP only and not the other nucleotides will be discussed in further detail in the later chapters along with supporting data. Possible future experiments supporting the proposed guanine oxidation as the cause for the observed change in quenching capability will be outlined briefly in the concluding remarks.

2 Sample Preparation and Experimental Method

2.1 Laser setup and TIR Illumination

A Nikon Eclipse TE-2000U™ inverted microscope with TIRF attachment was used for single-molecule detection using 2 HE-Neon Argon lasers from Melles Griot, California. The red laser optimized for illumination at 633nm, max. power output 10mW was used for Cy5 excitation with appropriate filter cube. The green HeNe laser at 543nm wavelength with an original max power 5mW was used for Cy3 excitation. Appropriate fluorescence filter cubes block any reflected and scattered laser light whilst still allowing the stokes’ shifted fluorescent light from the dye to pass through to the camera detector. A picture depicting the Microscope and cushioning air table with lasers underneath is shown on below.

Figure 13) Microscope with red (top left) and green (middle left) laser set-up on air- cushioned table.

Picture taken from lab showing Nikon TE2000U TIRF Microscope.

Below and on the following page is a diagram showing how the appropriate set of filter cubes, for both Cy3 and Cy5, work in allowing certain band-widths through for excitation and at same time still allowing different band-widths through for emission. Although the cubes depicted are not quite the same as the ones used in the experiment they are a close approximation and help visualize to the reader the importance of choosing the right filter cubes in combination with the laser to get as much fluorescence signal as possible and at same time trying to minimize noise from the any scattered and back reflected laser light.

Figure 14 a) Absorbance and emission spectral profiles for the cyanine dyes, Cy3 and Cy5, along with their appropriate filter cubes allowing maximum desired light through with minimal overlap of unwanted light.

Figure 14 b): continued

Above figures taken from http://www.microscopyu.com/articles/fluorescence/filtercubes/html

The camera used for detecting the emitted photons was an intensified CCD Photometrics cascade, back illuminated (512b) camera from Roper Scientific™ It had the required sensitivity to detect the weak signal from single molecules.

Obtaining total internal reflection fluorescence (TIRF) requires that the incident laser beam appears to come from near the edge of the objective lens before it hits the glass/sample solution interface and is reflected back inwards toward the other side of the objective again using appropriate lenses and mirror combinations. The objective used was a Plan Apo n=1.45, 60x objective from Nikon. Fluorescence immersion oil, type DF, was used with a refractive index of 1.52. The criteria for satisfying internal reflection at the glass/sample interface is that the light traveling between the 2 boundaries of different refractive index n1 and n2 will be bent away from the higher index medium, in this case glass (n1=1.52), towards the lower index medium, sample solution (n2=1.34). The critical angle required for TIR comes from Snell’s Law and is given by the following equation:

−1 n o θ ==sin2 , 61.8 eqn.[6] n1 The angle of incidence was adjusted by turning the vertical and horizontal laser alignment knobs on the TIRF attachment module.

The intensity of the surface illumination is a function of the incidence angle as well as the illumination depth, see figure 15. Thus one has to optimize the detection by adjusting the illumination close to the critical angle and not just by merely obtaining TIR. This process therefore needed some degree of fine tuning. There are few methods to adjust the illumination angle. For example by placing a prism on the objective and following the illumination direction focused some distance away. This is not quite the angle for evanescence as the angle from the tilted prism face is different than that from the flat sample direction. Another way that was used in this work is to use the reflected laser beam from the sample. The return beam from the objective is reflected by the dichroic beam splitter onto a wall a couple of meters away and its position is marked for a reference, once the optimum angle has been determined one can repeat this angle by returning to this mark. In order to determine the best angle, either the angle can be calculated empirically or by optimizing the signal from single molecule focused images.

Figure 15) Plot showing relative intensity of evanescent wave against displacement from surface, z (nm) and angle of incidence, θ (degrees) for 532nm wavelength excitation.

Graph slightly modified and taken from site, http://motility.york.ac.uk/poster/tirtheory.shtml

2.2 Cleaning Procedures and Surface Chemistry Protocols

One of the obvious requirements in achieving good reliable S-M images is of course to have as clean as possible sample surface. At first the coverslips (22mm sq x 0.17mm) were rinsed in pure water from Millipore® (0.22µm filtered) and then washed in isopropynol followed by another rinse in pure water. This simple process was enough to see fluorescence at higher concentrations i.e. 10’s of nano-Molar (nM) but was not good enough for S-M imaging which requires concentrations in the 10-25 pico-Molar (pM) range. Another problem was focusing onto the clean surface of the sample accurately enough to observe fluorescence the moment the laser was turned on. This is especially

40 important when observing Cy5 due to its much shorter bleaching lifetime time, <1s compared with Cy3, app. 10-20s. Lastly, in order to observe fluorescence for long periods of time a method for attaching the fluorophore-nucleotide-DNA to the surface securely whilst at same time still allowing the Cy3 molecule maximum freedom of movement was needed. These problems were addressed individually drawing on a lot of previous work [57, 81]. It should also be noted that the biotech company Helicos Biosciences, which supported this project, supplied the protocols for the surface chemistry and buffer preparation. These protocols were then adapted and modified for our experiments and sample preparation according to the project’s needs.

A simple and yet very effective way to focus much more easily on the glass slide surface was to make 2 or 3 light scratches, using a diamond scriber from VWR, on the middle of the slide only a few microns deep. This made it a lot easier to focus close to the surface and then only needing to make slight adjustments when it came to observing single-molecules attached to the slide. Even though the slides were scratched it was presumed that any adverse effect on the surface chemistry properties would be insignificant when focusing on an area away from the scratch.

2.2.1 Cleaning the glass slides

Once a batch of 16 slides had been gently scratched they were stored in 2 small coverslip mini-rack holders (C14784) from ‘Molecular Probes (invitrogen)’ and left to soak in 2ml/100ml liqui-nox® (Alconox Inc.) soap solution for several minutes then gently rinsed in pure water, 0.22µm filtered from Millipore™, and stored in a beaker containing more pure water. Next the slides were sonicated for about 20 minutes at room temperature before being treated using RCA protocol. They were bathed in a 4:6:1 mixture of Ammonia (NH4): H2O: Hydrogen Peroxide (H2O2) which was then left to boil gently for an hour in a fume cupboard. After being treated they were then rinsed again in pure water before being stored in another beaker containing filtered water. The slides

41 now have a slight negative charge imparted on them from the RCA protocol. This will help in the next procedure which is to build up layers of alternating charge using polyelectrolytes.

The main reason for building up layers of charge onto the glass surfaces is to facilitate a further chemical treatment procedure involving NHS-EDC, (N- Hydroxysuccinimide) - (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide), binding to the surface followed by attachment of the protein streptavidin. This then allows easier and more stable binding of the DNA template to the glass slide and reduces non-specific binding of any freely diffusing dNTP-Cy3/5 nucleotides in the sample solution. The fact that the last layer of charge built up on the slide is negative also helps repel the negatively charged phosphate backbone of the DNA template therefore helping to keep it upright and more rigid thus reducing unwanted steric effects on the fluorophore. This means that the attached fluorophores are less prone to be stuck to the surface or tangled up under the DNA/oligonucleotide lying on the surface itself. Under these conditions the fluorescence profile of the Cy3 dye would be biased due to its restricted ability to rotate more freely. This might cause the dipole to be oriented more favorably in one direction than another affecting the overall fluorescent intensity of the dye. A flexible more free dye is thus able to sample the full range of absorbance orientations and therefore any measurements observed are less prone to misinterpretation.

2.2.2 Application of Polyelectrolyte Multilayers

Polyacrylic acid (PACr) and Polyallamine (Pall) from Aldrich, negative and positively charged polymers respectively, were mixed at 750mg (40% by weight) and 300mg (2mg/ml) in 150ml each of pure water. They were then individually filtered through separate 0.22um filters. Both solutions were then brought to pH8.0 using NaOH and then stored in 2 wrapped clean beakers. To test if the mixtures were reactive several drops of both were added to a 1.5ml aliquot and a milky suspension was observed as a

42 positive test result for activity. [81] PACr and Pall solutions would keep their charge for several months. The slides were soaked first in the positive Pall for 10 minutes followed by rinsing and dipping in pure water for another 8 - 10 minutes and then soaking in the negative PACr for a further 10 minutes with a final rinse and soak in pure water for about 8 – 10 minutes once more. This constitutes one complete cycle of electrolyte layer addition and is repeated a further 3 times so as to build up enough negative charge on the final application.

Once the final layer of negative charge has been applied and the slides have been washed it is then necessary to go through the second surface chemistry protocol. This is taken from Helicos and modified for more practical use in our lab.

2.2.3 Surface activation for further attachments

NHS (N-Hydroxysuccinimide) (from Pierce lot#FJ73595) - EDC (1-ethyl-3-(3- dimethylaminopropyl) carbodiimide) (from Aldrich lot#124K1337) chemistry is a well used binding technique between liquid and solid surfaces used to activate certain types of functional end groups such as amine (NH2) groups or carboxyl (COOH) groups. In this case when the NHS-EDC is added to the slide surface in solution the surface becomes activated by the modification of the carboxymethyl groups to N-Hydroxysuccinimide esters [82]. In other words the COOH end groups become available on the surface as binding sites for the streptavidin.

Firstly two buffer solutions of approximately 600ml each were made up using MES (2-(N-Morpholino) ethanesulfonic acid) from Sigma (#M1317). One was 10 milliMolar (10mM) concentration of MES (6ml) at pH5.5 and the other a Coupling Buffer of 0.1M MES (60ml), 0.5M NaCl (17.32g) and brought to pH6.0. Both were diluted in pure water from Millipore™. The slides were then placed in 150ml of the 10mM MES buffer briefly before being transferred to another beaker of 0.22µm filtered 10mM MES and 10mM

EDC. They were left to incubate at room temp for an hour with continuous stirring using a small stir bar and rotating magnet plate device. After incubation the coverslips were again rinsed in another 150ml of 10mM MES buffer and then rinsed in 150ml of coupling buffer. A mixture of low molecular weight PACr (30ul or 0.2ml/L) with 10mM EDC and 150ml Coupling Buffer was made and the slides transferred to it and left to incubate with stirring for another 45 minutes. Again after incubation the slides were rinsed in coupling buffer this time and then stored in another beaker of coupling buffer and kept at 4deg C in fridge until further use.

2.2.4 Streptavidin Application Procedure

Streptavidin-plus® was bought from Prozyme® and is a widely used protein for binding between a surface and a biotinylated molecule such as DNA. It keeps the DNA immobilized to the surface therefore allowing long fluorescence observation with minimal disturbance and unwanted movement of the DNA oligonucleotides.

To add the streptavidin to the surface, a single glass slide was taken out from the coupling buffer previously and dried under a filtered air hose. In order to flow in and out different reagents and buffers hybriwell flow cells from GraceBiolabs were bought to place on top of the slides to form a sealed 18µl sample volume. These too were air dried and carefully set on top of the glass slide, scratched surface facing up. After gently pressing and sealing the hybriwell to the glass slide it was then possible to flow in and out the various solutions.

Coupling buffer was flushed in one hole by a pipette and out of the other hole in the flow cell using clean tissue, Kimwipes. This was possible by drawing out the solution through capillary action. Next approx. 4x50µl of 30mM NHS, 20mM EDC in coupling buffer was pipetted into the cell and left for 10-15 minutes to activate the surface. Any bubbles that formed were gently squeezed out to the edges of the volume area. More

44 coupling buffer was added to wash away any excess NHS-EDC from the surface before flowing in the streptavidin solution. The streptavidin was first diluted in coupling buffer to 10x concentration, 1.4mg/ml. Then 60 aliquots of 40µl each were made and stored at - 20ºC until needed with the excess diluted streptavidin kept in its original container. Each aliquot of 40µl after being diluted to desired 1x concentration of 0.14mg/ml in coupling buffer contained enough streptavidin for approximately two slides i.e. 200µl each. After flowing in the streptavidin and leaving it to bind to the surface for about 20 minutes 150- 200µl of coupling buffer was again washed in and out to get rid of any excess.

Figure 16) Diagram showing surface preparation prior to DNA attachment.

Starting from, a) RCA cleaned glass slide followed by addition of polyelectrolytes depositing alternative charged polymer tails onto surface, b). Following flowcell attachment to glass slide the surface is activated using an NHS-EDC reaction, c). Finally the binding protein streptavidin is added and attaches itself to the activated surface, d). Diagram modified and taken from Hebert and Braslavsky, "Single Molecule Fluorescence Microscopy and its Applications to Single Molecule Sequencing by Cyclic Synthesis," to be published in 2006 in, "New Sequencing Technologies" by Elsevier, edited by Keith Mitchelson.

2.2.5 Preparation and application of washing buffers

A few washing buffers have also been prepared. SSC buffer was used to clean the sample. It contains 0.45mol NaCl and 0.045mol of Saline Sodium Citrate and 0.1% by volume of TritonX-100 (a thick gel-like cleaning fluid). Approximately 5x75µl of SSC buffer was flowed into and out of the sample cell with the last wash cycle left to soak for about 10 minutes before pipetting in a PRB solution. This is a buffer solution with a more moderate pH 8.0 so that the DNA won’t be destroyed when added to the sample. It is made up of 20mM TRIS (pH 8), 50mM NaCl and 0.001% TritonX-100. All solutions and ingredients were purchased from Sigma/Sigma-Aldrich unless otherwise stated. Having now prepared the slide we are ready to add the fluorescently labeled DNA primer with template and the nucleotide/dye combination for observation under the microscope.

2.3 Properties of Cyanine Dyes, and DNA preparation

The cyanine dyes are known for having good photostability and high solubility in water which prevents clumping together especially at S-M level. They are relatively insensitive to changes in pH and show very strong ππ→ ∗ absorption [61] which thus presumably facilitates their interaction with the π -stack electron transfer pathway in the bases of DNA. Because they emit and absorb at different wavelengths they are a good combination to use where two distinguishable excitation wavelengths are required such as is the case in our experiment. The Cy5 tagged DNA primer is excited first in order to locate the DNA templates on the surface followed by excitation of the Cy3 labeled nucleotides to observe the amount and position of any incorporation onto the primer.

Figure 17) Chemical structure of Cy3-dCTP and Cy5-dUTP

Note the difference in length of the linker chain between the two heads of the Cyanine Dye. The 3 atom linker being Cy3 and 5 atom linker attributed to Cy5. It is the length of the linker chain along with the ∗ strong ππ→ absorption that is responsible for the various absorbance wavelengths from the visible, Cy3, to the far red/infra-red spectrum, Cy5 [61]. The other dNTP analogs are similar in structure. Picture taken from http://las.perkinelmer.com/content/TechnicalInfo/nel576.pdf .

The fluorescently labeled dNTPs were obtained from PerkinElmer Life Sciences, MA and were kept at -20ºC. To ensure reliable long term usage all fluorescent labels were stored in the dark and freeze-thaw cycles were kept to a minimum. All non-labeled dNTPs and the DNA polymerase, Klenow exo- (3’-5’) were purchased from New England Biolabs, NJ and kept under similar storage conditions. The main DNA template used was a single-stranded 128bp T404 with dual biotin at the 5’- end for attachment to the binding sites on the streptavidin coated surface. Another slightly different 128bp template, T407 with single biotin at the 5’- end was used in a second dUTP-Cy3 experiment to compare the quenching effects of an adjacent guanine base. A 30bp Cy3/Cy5, 5’-end labeled complementary primer sequence, P400 was then annealed to the template at its 3’-end. The dNTP-Cy3 analogs along with the help of the polymerase enzyme would then be incorporated onto the 3’- end of the primer and extend down the template strand one base at a time.

Figure 18) DNA templates, named T404 and T407 both with identical primers (in bold) shown underneath as complementary sequences growing from the 3’-end of the template, Oligos provided by Helicos Biosciences.

[T404] 5’Dual Biotin-A TCTCTCT G TCTCTCT A CTCTCTC G TCTCTCT A TCTCTCT G CTCTCTC A CTCTCTC G TCTCTCT ACTGCT ACTGCT ACTGCT ACTGCT ACTGCT ACT GCC CAC AAA CCA AAA GCC CAG ACA CCC GGA G-3’ 3’- - - - GG GTG TTTGGT TTT CGGGTCTGT GGG CCT C-5’-Cy5 [Cy5-P400]

[T407] 5’Biotin- A TCTCTCT G TCTCTCT A CTCTCTC G TCTCTCT A TCTCTCT G CTCTCTC A CTCTCTC G TCTCTCT CAG CAG CAG CAG CAG CAG CAG CAG CAG CAG CAG TCC CAC AAA CCA AAA GCC CAG ACA CCC GGA G-3’ 3’- - -GG GTG TTT GGTTTT CGG GTCTGT GGG CCT C-5’-Cy5 [Cy5-P400]

For T404 the first nucleotide to be incorporated will be dCTP-Cy3. The second experiment will require unlabeled dCTP and dATP-Cy3 as only the end nucleotide is to be fluorescent. Similarly the third experiment will have dCTP, dATP and dGTP-Cy3 and so forth for dUTP-Cy3. For the T407 template only one experiment was carried out using dATP, dCTP and dUTP-Cy3. These oligos where contributed by Helicos Biosciences.

5µl each of T404 and Cy5-P400 were added together and then diluted from 100µM down to 1µM in Tris buffer and clean water. The mixture was then heated to approx. 60ºC and allowed to cool slowly to let the primer anneal to its complementary fragment on the template. This was repeated for T407 and also for T404-Cy3-P400 giving three DNA-dye combinations in total. In the case of Cy3-P400-T404 where observation of the

48 labeled primer without addition of fluorescent dNTPs was desired the mixture was further diluted to concentrations suitable for S-M imaging i.e. 10-25pM. This was done so as to compare the bleaching rate of the Cy3-P400 primer by itself with the bleaching rates of the Cy3 incorporated nucleotides which were attached to the Cy5-P400 primer

A brief mention should be made about Fluorescence Resonance Energy Transfer (FRET) which is a process whereby an excited fluorophore such as Cy3 can transfer its energy to a suitable acceptor molecule such as Cy5 under the appropriate conditions. As both fluorophores are often used in FRET experiments in the 2-10nm scale it is possible though highly unlikely that some fluorescence will be quenched due to FRET interactions over the 30 base primer length of DNA. There have been cases of FRET occurring in oligonucleotides up to 35 bases in length [65] although the effect is minimal. One way to suppress FRET further is to bleach the acceptor fluorophore i.e. the Cy5-primer, before acquiring the movie of dNTP-Cy3 and this was done when using the red laser to focus on the DNA templates bound to the surface. It must be noted however that the possibility of some quenching due to FRET cannot be ruled out entirely but it is thought that this quenching is negligible compared to the overall quenching effect of the covalently bonded nucleotides.

The dNTP-Cy3 analogs were diluted in 10mM Tris buffer and pure water from their original concentration, 1mM, to 4µM and kept in 1.5ml aliquots at -20ºC until needed. Unlabeled dNTPs were also diluted to 4µM and refrigerated. The next nucleotide to be incorporated onto the T404-Cy5-P400 (T4P4Cy5) strand opposite the base guanine is of course dCTP and in the case of our experiment, dCTP-Cy3. In order for incorporation of the nucleotides to be successful it was necessary to use higher than S-M level concentrations. Otherwise the mixture would be too dilute and therefore a single- strandand nucleotide would have a very low probability of coming into contact. A concentration of 100nM T4P4Cy5 (10 µl) with approx. 300nM dNTP-Cy3 (7.5µl) in a final volume of 100µl gave the most incorporation. In addition, 10µl of a reaction buffer

and 1-1.25µl of Klenow exo- (50-65 units/ml) were added along with pure water to make the final 100µl volume. The aliquot was gently stirred and the polymerase enzyme was given 1 to 2 minutes to incorporate the dCTP-Cy3 nucleotides. The solution was then heated and allowed to cool back down to room temperature. Once cooled the solution was then diluted a couple of times in 10mM Tris and pure water to a concentration of 25pM Cy5 DNA/Primer and 75pM dCTP-Cy3.

Next, the DNA was applied to the surface. For an illumination area of approx. 40µm x 40µm one would need several hundred stable bound T404/Cy5-primer strands in order to have enough sequenced nucleotides to be statistically acceptable. By taking 25pMolar concentration of DNA-Cy5 and multiplying by avogadro’s number, 6.0x10²³ then dividing by 1 million one can figure out how many molecules there are per µl of sample solution. In this case approx.15 million molecules per µl. If one assumes a sample volume of 20µl then there are roughly 300 million molecules in the sample volume. Even if only one third bind to the streptavidin coated surface that still leaves 100 million templates bound in an area of approx. 1cm² or 10000µm x 10000µm. This equates to approximately 1 molecule per µm². As each pixel is 100nm x 100nm then every molecule has 100 pixels to itself on average. A 128bp strand of DNA is about 35nm long therefore it has a radius of about 15 times its own length before coming into contact with another strand. The negative charge on the glass surface tends to repel the negatively charged DNA backbone therefore helping to keep the template away from the surface except the connected tail.

Figure 19) DNA with incorporated nucleotides attached to glass surface.

Cy3dNTPs are attached to the last base on the 3’ end of the Cy5-primer sequence with the aid of klenow exo- polymerase. Purple squares at base of DNA template represent surface attachment chemistry from figure 16). Diagram modified and taken from Hebert and Braslavsky, "Single Molecule Fluorescence Microscopy and its Applications to Single Molecule Sequencing by Cyclic Synthesis," to be published in 2006 in, "New Sequencing Technologies" by Elsevier, edited by Keith Mitchelson.

The sample bin size of the fluorescent spot from a single-molecule in this experiment is 2-3 pixels in diameter so for a surface density of 1 molecule every 1µm² there is still plenty of room to resolve 2 molecules separately. With a possible 1600 molecules per illumination screen there should be more than enough incorporated nucleotides for observation. Of course other considerations should be taken into account. Variations in dye/DNA integrity, aging and degradation of previously prepared solutions and the ever present discrepancies between the preparation of one sample and the next lead to an inevitable distribution in sample quality and therefore the amount of nucleotide

incorporation present. The procedure for making the other dNTP-Cy3 and DNA/Cy5- primer samples are similar except that as each extra Cy3-nucleotide is being added along the 3’-end of the primer a non-labeled nucleotide must also be added to fill in where the previous dNTP-Cy3 nucleotide was incorporated. Now that the dye/DNA solution has been made 20-50µl are then pipetted into a previously prepared flow cell containing PRB and streptavadin coating. The sample is now ready for observation under the microscope and data acquisition using the camera detector and software.

2.4 Observation of sample and data acquisition

Imaging of the surface with the microscope through the camera was facilitated by the manufacturer’s imaging software and camera control, WinView program. The flow cell was first checked under the microscope for any dirt or miscellaneous dye on the surface before adding the 20-50µl dye/DNA/nucleotide solution. By focusing on the scratches at the surface through the eyepiece using ordinary white light and then obtaining the image on the computer screen through the camera we can visually adjust the image to get as close to the surface as possible. One can focus even closer to the surface by picking a part of the scratch that tapers off to a hair-like fine point and focus on the tip at the end. First, using the red laser and adjusting the camera data acquisition time to about 0.5 seconds per frame check for any dirt on the sample surface. Then adjust the TIRF attachment and filter cubes for the green laser and check for any excess dirt and dye.

Note: due to the relatively low green laser intensity and the lower extinction co- efficient of Cy3 compared to Cy5 it is necessary to increase the acquisition time per frame for Cy3 to about 2 seconds. This is in order to have similar intensities on screen for both dyes which will aid the correlation software, described in section 2.5, in comparing the two separate movies later.

After checking the hybriwell flow-cell is clean, 20-50µl of sample solution is pipetted in and any excess drawn out. The sample is then left for a few minutes to allow the DNA to bind to the streptavidin on the surface. 1-200µl of PRB solution is then washed in and out to get rid of any floating unbound DNA/nucleotides. This will further help reduce any background fluorescence. After focusing again on the surface as before we are now ready to fine tune the focus even closer. To do this a simple and yet extremely effective method was developed. By taking apart the original Nikon laser ‘TIRF’ illumination module that contains the filters and the pinhole which adjusts the sample illumination area and lining them up manually we were able to add a second adjustable pinhole aperture behind the first and slightly off center to it. Now it was possible to focus on a small corner of the sample area on screen without actually having the laser illuminating and thus bleaching the main part of the sample. This meant that when turning on the laser and acquiring the images of the rest of the sample we would immediately be in near perfect focus on the surface of the Cy5DNA-Primer strand without having to make any further adjustments. This method that is not widely used for TIRFM at the S-M level will greatly improve the ability of the experimenter to have high quality images the instant the laser and camera are turned on and thus no loss of bleached molecules when trying to focus on the sample otherwise.

Figure 20) Separated Laser and filter module with 2 pinhole apertures.

Fiber optic cable enters from left to collimator with incident angle adjustment knobs. The beam then passes through the filter block (middle) and into the back of the microscope via 2 pinhole apertures. 1st one is fixed in position and second one can be moved in and out as necessary. Picture taken from Laboratory.

The cascade camera has on chip multiplication gain which allows it to greatly increase the signal received from relatively few photons. The gain was tunable from 0 (no gain) to a maximum of 4095. This scale is based on an exponential-like relationship so a small increase in gain greatly magnifies the number of electrons cascading through the cameras detection system to boost the signal. It was not necessarily better to have maximum gain on as the background noise level also gets increased therefore a compromise between signal and background noise was found.

To acquire images of the DNA/primer-Cy5 the gain of the camera was set at app. 2500 with an acquisition time of 0.5 s. The illumination area was adjusted to 40µm x 40µm using the main pinhole. Due to the much shorter bleaching life-time of Cy5 compared to Cy3 most of the Cy5 was bleached after only a few seconds. This was perfectly acceptable as only the first couple of frames were needed showing all of the Cy5-primer/DNA bound molecules.

After re-adjusting the microscope for use with the green laser it was now possible to acquire images of the attached Cy3-dNTP molecules. The gain was set at around 3000 with an acquisition time of 2 seconds. The bleaching life-time for Cy3 is much longer than Cy5 and, coupled with the much reduced power output of the laser it became even more so. On average it took around 500-600 frames of 2 seconds each to get enough molecules bleached within the desired sample area.

2.5 Initial data analysis using custom developed correlation software

Upon acquiring movies of both red and green laser illumination it is now necessary to crop them together and check for the amount of, if any, correlation between incorporated nucleotides and the DNA template. This was done using pre-developed software packages, correlator.exe by Helicos and a suite of programs written in IDL (an imaging software platform) that have been developed by Hebert and Braslavsky based on Crocker object tracking programs [57], [83] With these programs it is possible to crop together movies of both red and green lasers and correlate them. The program analyzes each pixel and takes any objects that fit the parameters for S-M e.g. above a certain threshold background intensity and stores their position in each frame. Every subsequent frame is then compared to the first one and the more overlap between selected objects in each frame the higher the degree of correlation.

Figure 21) Showing amount of correlation from 1st movie frame using the red laser with all subsequent frames using the green laser.

Figure 21 a) shows the actual amount of correlation of Cy3 with Cy5. A central peak is indication of good correlation between movies i.e. lots of Cy3-dNTPs incorporated onto the Cy5DNA strand therefore minimal change of position from one frame to the next. Figs. b) and c) show 1st frames of movie under red and green laser respectively. Picture taken from actual dCTP-Cy3 data.

After many attempts and different combinations of DNA/nucleotide concentrations along with choosing appropriate acquisition times and camera gain levels as well as adjusting various other parameters to maximize correlation of nucleotides onto their DNA primers it was possible to obtain several long movies of all four DNA-Cy5/dNTP- Cy3 configurations along with the DNA-Cy3 primer by itself.

3 Results and comments

3.1 Editing and analysis of movie data

Once the data acquisition process has been completed using the Winview software package, the correlator.exe program is used to analyze the alignment of the images between the red and green illumination and in between the frames and then to put together a movie of the desired sample area under illumination. The fact that the laser illumination is not uniform over the whole 512x512 pixel area but rather has a gaussian like distribution means a spot on the edge of the movie area will have a lower intensity compared to a spot in the middle. By choosing a smaller movie area near the center we can minimize variation of background and peak intensities that are not due to the particular dNTP-Cy3 combination itself. Although it would have been more desirable to have every movie exactly the same size it soon became apparent that some movies would have to be larger than others due to lesser numbers of incorporation events in certain samples. The most important criteria that had to be satisfied was to have enough single- molecules that bleached within a certain amount of time so that the statistical sample size was large enough (app.100 incorporated nucleotides) to approximate ensemble measurements. This meant that some movie areas were larger than others to ensure roughly the same amount of single-molecules per movie. The smallest movie size was 150x150 pixels and the largest was 250x250 pixels. Although this seems like a large difference, the variation in background intensity compared to the whole 512x512 pixel area was small.

Of interest to note was that the average background intensity at the start of a 100 frame movie clip was lower compared to the last 100 frames. This increase in the overall background fluorescence is not clear but might be due to damage on the surface by radical molecules produced in the experiment, but the variation in background intensity over the sample area within each frame is minimal and this gradual level of background increase still allowed for data analysis to be performed.

Once the final movie was put together for each configuration it was then analyzed using the IDL based software. We developed some further sub-routines in IDL to analyze each individual S-M trace within a movie and choose the traces that were most likely to be incorporated nucleotides. After going through all the S-M traces and summing up the overall intensities of the incorporated ones, graphs were obtained for each dNTP-Cy3 setup showing the exponential bleaching behavior as expected at the ensemble level [6, 65, 75, 76, 84]. Further analysis of the data has been done with microsoft Excel and converted into graphs from there.

The summed up intensity profiles for each dNTP-Cy3 combination were fitted with several curves to ascertain the best fit. A bi-exponential fit, as was shown earlier in chapter 1 along with a mono-exponential fit with and without constants were all

2 ()EOxpbs− attributed to each data set and their minimized χ 2 values, defined as ∑ Exp were compared. A graph of each dNTP-Cy3 setup with normalized fluorescence intensity versus time is shown along with selected S-M traces highlighting typical individual profiles. A selection of frames from each original movie acquisition is also shown revealing the overall bleaching pattern of many molecules followed by a brief discussion on the behavior of each dNTP-Cy3/DNA analog. Special emphasis will be given to the fluorescent profiles of dGTP-Cy3/DNA and their possible evidence for guanine oxidation.

3.2 Cy3-Primer/DNA by itself.

Next I will show all intensity data obtained from the single molecules analyzed. The Cy3-Primer/DNA was used as an extra configuration to compare the bleaching lifetimes with just one fluorophore attached to the DNA and no dNTP-Cy3 incorporated at the 3’- end of the primer sequence.

Figure 22) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of Cy3-DNA/Primer.

Fluorescence Intensity Profile of Cy3-Primer

0.9

0.8 data bi-exp+C 0.7 1exp+C 0.6 bi-exp 1exp 0.5 0.4

Relative Intensity 0.3 0.2 0.1 0 0 50 100 150 200 250 Time (seconds)

Figure 22): continued

a) Single-Molecule Traces Cy3P4T4 b) Single-Molecule Traces Cy3P4T4

200 250 S-M# 1 S-M#111 150 S-M#26 200 S-M#153 S-M#79 150 100 S-M#204 100 50 50 Intensity (a.u) (a.u) Intensity Intensity (a.u) 0 0 0 50 100 150 200 250 0 50 100 150 200 250 Time (seconds) Time (seconds)

Single-Molecule Traces Cy3P4T4 c) Graph shows the total ‘normalized to maximum fluorescence’ intensity vs time of all bleached 150 S-M#280 single-molecules of Cy3-Primer/T404. Mono- 100 S-M#300 and bi-exponential curves were fitted using a S-M#325 least squares fit analysis with chi-squared 50 values of 0.248, 0.248, 0.337 and 0.343 for bi- exp+c, bi-exp, 1exp+c and 1exp respectively. Intensity (a.u) (a.u) Intensity 0 Plots a-c, show absolute intensities vs time, of 9 0 50 100 150 200 250 selected S-Ms of Cy3-Primer/DNA and their Time (seconds) sudden bleaching steps.

The decrease in fluorescence is described sufficiently well by a mono-exponential fit with no significant improvement using a bi-exponential curve. The bleaching life- τ time, b , is app. 120 seconds and a total of 132 S-Ms were analyzed out of over 350 objects with only about 7 or app. 5% being completely quenched at the onset of illumination.

Shown on next page are selected frames from the acquired movie of the Cy3- Primer/DNA showing the decrease in number of bright ‘spots’ (S-Ms) as the number of photobleached molecules increases with time.

Figure 23) Selected frames from Cy3-Primer/DNA movie illustrating bleaching behavior over time.

200x200 pixel (20um x 20um) Cy3Primer/DNA movie (125frames x 2secs each) showing from top left to bottom right, frames 1, 25, 50, 75, 100 and 125.

3.3 dATP-Cy3/DNA.

Figure 24) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dATP-Cy3-DNA.

(a) Fluorescence intensity profile of dATP-Cy3/DNA

0.6 data 0.5 Bi-exp+C 0.4 1exp+C Bi-exp 1exp 0.3 0.2

Relative Intensity 0.1

0 0 200 400 600 800 1000 1200 Time (seconds)

Single-Molecule Fluorescence Traces Single-Molecule Fluorescence Traces dAtpCy3/DNA_Aug dAtpCy3/DNA_Aug

150 150 S-M#22 S-M#103 S-M#73 100 100 S-M#183 S-M#82 S-M#213 50 50

(a.u) Intensity 0 (a.u) Intensity 0 0 200 400 600 800 1000 1200 0 200 400 600 800 1000 1200 b) Time (seconds) c) Time (seconds)

Single-Molecule Fluorescence Traces Graph ‘a’ shows the total ‘normalized to dAtpCy3/DNA_Aug maximum fluorescence’ intensity vs time of all

100 S-M#292 bleached single-molecules of dATP-Cy3/DNA. 80 S-M#350 Mono-and bi-exponential curves were fitted using 60 S-M#366 a least squares fit analysis with chi-squared values 40 of 0.731, 0.734, 0.734 and 1.811 for bi-exp+C, bi- 20 exp, 1exp+C and 1exp respectively.

Intensity (a.u) (a.u) Intensity 0 Plots ‘b-d,’ show absolute intensities vs time, of 9 0 200 400 600 800 1000 1200 typical individual S-M traces of dATP-Cy3/DNA d) Time (seconds) and their sudden bleaching steps.

Figure 25) Selected frames from dATP-Cy3/DNA movie showing decrease in Fluorescent S-Ms due to Photobleaching.

From top left; frame 1 showing Cy5-Primer/DNA locations under red laser. Frames 5, 100, 225, 350, 475 and 600 show dATP-Cy3 positions either incorporated into the DNA- Primer or not. Notice not all the molecules were bleached but only those that did bleach were analyzed and their fluorescence profiles added up to obtain the graph in Fig.24 (a). Scale bar shown is 5µm. Movie area is 225 x 225 pixels or 22.5µm x 22.5µm.

There was very little difference in the chi-squared fit values for the bi-exponential curves and the mono-exponential + constant curve. The much larger χ 2 value in the single exponential curve with no constant can be explained by the lack of a term to account for the residual background influence. As can be seen from graph ‘a’ the fluorescence does not decay to zero which would be expected because all the selected molecules contributing to the graph were bleached. This non-zero base value of 0.057 therefore implies that there is some background influence. Also, the fact that the second exponential term in the ‘bi-exp’ curve and the constant term in the ‘1exp+C’ curve are both almost identical (i.e. 0.057) shows this to be the case. The bleaching life-time for τ dATP-Cy3/DNA, b , is app. 360 seconds which is appropriate for the low illumination intensities used from a much weakened laser.

There were 131 single-molecules incorporated that were bleached in the sample out of almost 400 objects analyzed by the correlator software. Of these, only around 6 molecules or <5% were completely quenched at the start of illumination.

3.4 dCTP-Cy3/DNA.

Figure 26) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dCTP-Cy3-DNA.

(a) Fluorescence intensity profile of dCTP-Cy3/DNA

0.5 data 0.4 bi-exp+C 1exp+C 0.3 Bi-exp 1exp 0.2

Relative Intensity Intensity Relative 0.1 0 0 200 400 600 800 Time (seconds)

Single-Molecule Traces dCtpCy3/DNA Single-Molecule Traces dCtpCy3/DNA 150 250

S-M# 6 S-M#200 S-M#36 200 S-M#216 (2 100 S-M#84 150 mols) S-M#249 50 100

(a.u) Intensity 50 Intensity (a.u) (a.u) Intensity 0 0 0 200 400 600 800 0 200 400 600 800 b) Time (seconds) c) Time (seconds)

Graph ‘ a’ shows the total ‘normalized to Single-Molecule Traces dCtpCy3/DNA maximum fluorescence’ intensity vs time of all 250 bleached single-molecules of dCTP-Cy3/DNA. S-M#305 200 S-M#312 Mono-and bi-exponential curves were fitted 150 S-M#371 using a least squares fit analysis with chi-squared 100 S-M#396 values of 0.81, 0.81, 0.87 and 1.07 for bi-exp+C, 50 bi-exp, 1exp+C and 1exp respectively. Intensity (a.u) Intensity 0 Plots ‘b-d,’ show absolute intensities vs time, of 0 200 400 600 800 9 typical individual S-M traces of dCTP- d) Time (seconds) Cy3/DNA and their sudden bleaching steps.

Once again the decrease in fluorescence due to photobleaching is sufficiently τ described by a single-exponential equation. The values for the bleaching lifetime, b , however, varied considerably between fits, ranging from approx. 520 seconds for the bi- ττ+ 12 exp+C, , 925 seconds for 1exp+C, 336 seconds for a bi-exp fit and finally 544 2 seconds for 1exp. From the graph it can be seen that the bleaching lifetime is approximately 550 seconds which would indicate that the 1exp curve gives a more than adequate explanation of the fluorescence profile. 145 bleached S-Ms were analyzed out of just over 400 objects correlated and only 7 of these or <5% were completely or partially quenched from the start. This is similar to dATP-Cy3/DNA except for the over 50% longer bleaching lifetime of dCTP-Cy3/DNA.

Figure 27) Selected frames from dCTPCy3/DNA movie showing decrease in fluorescent S-Ms due to photobleaching.

Figure 27): continued

From left to right: Frame 1 showing Cy5-Primer/DNA locations under red laser. Frames 2, 100, 200, 300, 400 and 500 show dCTP-Cy3 positions either incorporated into the DNA- Primer or not. Again, not all the molecules were bleached but only those that did bleach were analyzed and their fluorescence profiles summed up to obtain the graph in Fig.26 (a). Scale bar shown is 5µm and sample area is 150 x 150 pixels or 15µm x 15µm.

3.5 dUTP-Cy3/DNA (T404).

Figure 28) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dUTP-Cy3/DNA.

(a) Fluorescence intensity profile of dUTP-Cy3/DNA

0.6

data

0.5 bi-exp+C 0.4 1exp+C bi-exp 1exp 0.3 0.2

Intensity Relative 0.1 0 0 200 400 600 800 1000

Time (seconds)

Figure 28): continued:

Single-Molecule Traces dUtpCy3/DNA Single-Molecule Traces dUtpCy3/DNA

120 200

S-M#26 S-M# 82 150 80 S-M#34 S-M#124 S-M#49 100 S-M#153 40 50 ntensity (a.u) ntensity ntensity (a.u) ntensity I 0 I 0 :0200400 600 800 1000 0 200 400 600 800 1000 b) Time (seconds) c) Time (seconds)

Single-Molecule Traces dUtpCy3/DNA Graph ‘a’ shows the total ‘normalized to maximum fluorescence’ intensity v time of all 200 bleached single-molecules of dUTP-Cy3/DNA. S-M#186 150 Mono-and bi-exponential curves were fitted S-M#200 using a least squares fit analysis with chi- S-M#218 100 squared values of 1.07, 1.34, 1.88 and 1.79 for 50 bi-exp+C, 1exp+C, bi-exp and 1exp ntensity (a.u) ntensity

I respectively. 0 Plots ‘b-d,’ show absolute intensities v time, of 0200 400 600 800 1000 9 typical individual S-M traces of dUTP- d) Time (seconds) Cy3/DNA and their sudden bleaching steps.

As one can see from Fig. 28(a) there is a noticeable increase in the relative fluorescence intensity within the first 40-50 frames of the movie. It is not clear why this occurs as dUTP is the least oxidizable of the four nucleotides and if anything would be more likely to bleach faster due to supposedly less quenching. A simple explanation might arise from adjustments made to focusing of the laser during a movie although upon examination of the background fluorescence it was found that there was hardly any unusual increase in intensity other than the normal, slight, gradual increase over a long period of illumination in a 10 minute movie.

Figure 29) Selected frames from dUtpCy3/DNA movie showing decrease in fluorescent S-Ms due to photobleaching.

From left to right: Frame 1 showing Cy5-Primer/DNA locations under red laser. Frames 10, 100, 200, 300, 400 and 500 show dUTP-Cy3 positions either incorporated into the DNA-Primer or not. Again, not all the molecules were bleached but only those that did bleach were analyzed and their fluorescence profiles summed up to obtain the graph in Fig.28 (a). Scale bar shown is 5µm and sample area is 225 x 225 pixels or 22.5µm x 22.5µm.

Only 82 S-M traces with incorporated nucleotides were bleached and analyzed in this sample out of app.250 objects. About 27 or >30% of these showed partial quenching of fluorescence though very few were completely quenched. Nearly all of the 27 partially quenched molecules showed an increase in fluorescence around the 20th frame or 40 seconds into the movie. This can be seen on the overall fluorescence in graph 28 (a). There is also the fact that more than 1 dUTP-Cy3 molecule could have been sequenced on the strand as all 4 nucleotides were washed in and out in this particular experiment. The fact that most of the quenched molecules increased their fluorescence at practically the same time almost certainly points to an unforeseen experimental error occurring around that frame time rather than perhaps some intrinsic property of dUTP-Cy3 itself.

On the other hand if one takes a look at the extended primer sequence in dUTP- Cy3/DNA (T404) it is clear that the adjacent previous base to the labeled uracil nucleotide is in fact guanine. It is possible that the nearby guanine base has an extended quenching effect on the dUTP-Cy3 molecule. It has been shown [53, 85] that nearby guanine bases and doublets especially to the 5’-end can quench fluorescence of nearby molecules on the same strand by as much as 40%. These guanine sites act as traps or holes for radicals to migrate down the ‘pi’-stack of the DNA strand. It is possible that a guanine base can alter the oxidation potential of uracil and therefore dUTP-Cy3 thus perhaps increasing its quenching capabilities until the guanine reacts with a radical. Unfortunately using this explanation would imply that dCTP-Cy3’s fluorescence should also show an increase as it has 3 guanines to its immediate 5’-end. However, it must be said that each combination has a different effect on various base sequences because they all have their own specific redox potentials which are altered slightly due to coupling effects from e.g. hydrogen bonds between base pairs and de-localization of the ‘pi’-stack molecular orbitals from the surrounding intrastrand bases [18, 50, 72, 86].

In other words a simple base by itself will have a certain redox potential which will differ compared to its nucleotide derivative which will change again upon incorporation into single and double stranded DNA. Usually the net effect is to lower the oxidation potential when going from single base to nucleotide etc. [35] In order to help alleviate the ambiguity in analyzing the dUTP-Cy3/DNA fluorescence another experiment was carried out on dUTP-Cy3 except this time sequenced to the T407- DNA template. The primer sequence was the same as before.

3.5.1 dUTP-Cy3/DNA (T407).

Figure 30) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dUTP-Cy3/DNA (T407).

(a) Fluorescence intensity profile of dUTP-Cy3P4T7

0.7 data 0.6 Bi-exp+C

1exp+C 0.5 Bi-exp 1exp 0.4 Intensity 0.3 0.2

Relative 0.1

0 0 200 400 600 800 1000 1200 Time (seconds)

Figure 30): continued

Single-Molecule Traces dUtpCy3P4T7 Single-Molecule Traces dUtpCy3P4T7

400 800 S-M#70 300 S-M# 7 600 S-M#80 200 S-M#33 S-M#109 S-M#42 400 100 200 Intensity (a.u) (a.u) Intensity (a.u) Intensity 0 0 0 200 400 600 800 1000 1200 0 200 400 600 800 1000 1200 Time (seconds) c) b) Time (seconds)

Graph ‘a’ shows the total ‘normalized to maximum fluorescence’ intensity vs time of all Single-Molecule Traces dUtpCy3P4T7 bleached single-molecules of dUTP-Cy3/DNA 400 (T407). Mono-and bi-exponential curves were 300 S-M#161 fitted using a least squares fit analysis with chi- S-M#175 squared values of 1.07, 1.34, 1.88 and 1.79 for 200 S-M#189 bi-exp+C, 1exp+C, bi-exp and 1exp 100

Intensity (a.u) (a.u) Intensity respectively. 0 Plots ‘b-d,’ show absolute intensities vs time, of 0 200 400 600 800 1000 9 typical individual S-M traces of dUTP- d) Time (seconds) Cy3/DNA (T407) and their sudden bleaching steps.

For dUTP-Cy3/DNA (T407) there was a much smaller initial rise in relative fluorescent intensity compared to dUTP-Cy3/DNA (T404). This shows that there were perhaps some strands with more than 1 fluorophore sequenced on the previous dUTP- Cy3/DNA (T404) experiment. Looking at S-M#’s 189 and 161 from figure 30 (d) one can see complete quenching of the fluorescence both at the start of laser illumination as in S-M# 189 until app. 475 seconds after laser has been on and also, in the case of S-M# 161 complete quenching in between periods of fluorescence.

Figure 31) Selected frames from dUTP-Cy3/DNA (T407) movie showing decrease in fluorescent S-Ms due to photobleaching.

From left to right: Frame 1 showing Cy5-Primer/ DNA (T407) locations under red laser. Frames 4, 100, 225, 350, 475 and 600 show dUTP-Cy3 positions either incorporated into the DNA-Primer or not. Again, not all the molecules were bleached but only those that did bleach were analyzed and their fluorescence profiles summed up to obtain the graph in Fig.30 (a). Scale bar shown is 5µm and sample area is 150 x 150 pixels or 15µm x 15µm.

Even though the sample area was only 150 x 150 pixels a relatively high number of molecules were incorporated and analyzed. Out of just about 250 objects in the sample area 108 were valid bleached S-Ms. Thus over 40% of molecules analyzed by the software contributed their fluorescence profiles to graph 30 (a). This can be explained by the fact that the T407 template was only used in one experiment and so when annealed to the Cy5-Primer the mixture was ‘fresh’ and had been in and out of the fridge much less than the Cy5-Primer T404 sample. Thus it would have undergone less freeze thaw cycles which are prone to degrading the quality of the DNA and dye sample over time.

Of the 108 S-Ms chosen about 13 or 12% exhibit strong quenching from the start. This is less than half compared to the dUTP-Cy3/DNA (T404) conjugate. The absolute intensities of individual molecule traces of the dUTP-Cy3/DNA (T407) were noticeably higher than the other configurations and this again could be due to the samples being fresher than their T404 counterparts with the dyes spending much less time in the light.

The fluorescent profiles also seemed to be much more erratic than the other samples with many showing more than one distinguished peak. It is possible that the Cy3 dye aligns itself in specific orientations due to solvent-dipole interactions and instead of rotating ‘freely’ is fixed in one favored position for an extended period of time before eventually re-orientating itself. It is still not clear why the erratic peeks and plateaus in fluorescence appear to be more common in dUTP-Cy3 than the others.

3.6 dGTP-Cy3/DNA.

Figure 32) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dGTP-Cy3/DNA

(a) Fluorescence intensity profile of dGTP-Cy3/DNA 0.5 data 0.4 bi-exp+C 1exp+C 0.3 bi-exp 1exp 0.2 0.1 Relative Intensity Intensity Relative 0 0 200 400 600 800 1000 Time (seconds)

- Single-Molecule Fluorescence Traces SingleMolecule Fluorescence Traces dGtpCy3/DNA dGtpCy3/DNA

250 250 S-M#78 200 200 S-M#10 S-M#97 (a.u) S-M#35

( 150 150 S-M#124 S-M#54 100 100 50 50

Intensity (a.u) 0 0 Intensity 0 200 400 600 800 b) 0 200 400 600 800 Time (seconds) c) Time (seconds)

Single-Molecule Fluorescence Traces Graph ‘a’ shows the total ‘normalized to dGtpCy3/DNA maximum fluorescence’ intensity vs time of all 160 bleached single-molecules of dGTP-Cy3/DNA. S-M#199 120 S-M#225 Mono-and bi-exponential curves were fitted using a least squares fit analysis with chi-squared values 80 S-M#262 of 0.92, 22.2(!), 4.80 and 5.05 for bi-exp+C, bi- 40 exp, 1exp+C and 1exp respectively.

Intensity (a.u) Intensity 0 Plots ‘b-d,’ show absolute intensities vs time, of 9 d) 0 200 400 600 800 typical individual S-M traces of dGTP-Cy3/DNA Time (seconds) and their sudden bleaching steps.

One can already clearly see the marked difference in the shape of the fluorescence profiles of dGtpCy3/DNA compared with the other nucleotide/DNA combinations. It is obvious from looking at figure 32 (a) that the fluorescence intensity is not best described by a single-exponential function but rather is much better described by a bi-exponential decay relation. All 9 S-M traces shown exhibit either complete or severe fluorescence quenching at the start of illumination with most being quenched between 50 and 150 seconds before starting to fluoresce and finally bleach after continued illumination under the laser. Out of the 99 molecules contributing to the graph over 90 showed complete initial quenching of Cy3 fluorescence. With the ambiguous exception of dUTP, this corresponds to over 16 times the number of fluorescent single-molecules being totally quenched compared with the other nucleotide configurations. The two bleaching lifetimes τ τ 1 and 2 of dGTP-Cy3/DNA for the bi-exponential fits were 106 and 113 seconds for the bi-exp+C curve and 154 and 162 seconds for the bi-exp fit. For the 1exp+C and 1exp τ χ 2 curves, b was 342 and 457 seconds respectively. The exceptionally large value of 22 for the bi-exp fit compared to 0.92 for the biexp+C could again be explained by the lack of a constant term to account for the background intensity. From observing the last frame in the dGTP-Cy3 movie one can see that there are hardly any unbleached molecules left therefore the constant background intensity dominates and does not fit an exponential decay relation. One can also see from the graph that the tail end of the data line has almost leveled out indicating that a ‘constant’ term relating to the background fluorescence will be the dominant contributor. The bi-exp fit tends to diverge from the data at the tail end and thus helps to explain the relatively large χ 2 value.

Several other dGTP-Cy3 movies were recorded that showed the same increase in S- Ms fluorescing after some time and one was chosen for further analysis. This was done in order to show that the initial rise in fluorescent molecules was not due to some sort of inherent error in the experimental method or equipment or perhaps because of a bad sample.

Figure 33) Selected frames from dGTP-Cy3/DNA movie showing decrease in fluorescent S-Ms due to photobleaching.

From left to right: Frame 1 showing Cy5-Primer/DNA locations under red laser. Frames 5, 75, 125, 200, 350, and 500 show dGTP-Cy3 positions, using the green laser, either incorporated into the DNA-Primer or not. This time almost all the molecules were bleached and those that were incorporated into the DNA were analyzed and their fluorescence profiles summed up to obtain the graph in Fig.32 (a). Scale bar shown is 5µm and sample area is 150 x 150 pixels or 15µm x 15µm.

NOTE: After observing the Cy5-Primer in frame 1 one can see a drastic drop in the number of fluorescing dGTP-Cy3/DNA molecules in frame 5. This corresponds to guanine’s superior quenching characteristics compared with the other nucleotides. After some time one can see an increase in the number of fluorescent single molecules, frame 75, followed by the inevitable decrease in the number of fluorescent S-Ms due to photobleaching under continued illumination.

A discussion outlining possibilities for interpreting this onset of fluorescence, after temporary complete quenching, as guanine oxidation will be given in section 4) after the following supporting data from the second dGTP-Cy3/DNA sample.

3.6.1 dGTP-Cy3/DNA (sample 2)

Figure 34) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dGTP-Cy3/DNA (sample 2).

(a) Fluorescence intensity profile of dGTP-Cy3/DNA (#2)

0.3 Data 0.3 Bi-exp+c 0.25 1exp+c

Intensity Intensity Bi-exp 0.2 1exp 0.15 0.1 Relative 0.05 0 0 250 500 750 1000 1150 Time (seconds)

Single-Molecule Traces dGtpCy3/DNA_sept Single-Molecule Traces dGtpCy3/DNA_sept

400

S-M#137 300 S-M#15 S-M#114 S-M#13 300 200 S-M#157 S-M#32 200 100 100 Intensity (a.u) Intensity

Intensity (a.u)) 0 0 0 250 500 750 1000 0 200 400 600 800 1000 1200 Time (seconds) Time (seconds) b) c)

Figure 34): continued

Single-Molecule Traces dGtpCy3_sept Graph ‘a’ shows the total ‘normalized to maximum fluorescence’ intensity vs time of all 300 bleached single-molecules of dGTP-Cy3/DNA S-M#212 (#2). Bi-and mono-exponential curves were fitted 250 S-M#261 200 S-M#272 using a least squares fit analysis with chi-squared 150 values of 0.84, 0.95, 2.55 and 4.41 for bi-exp+C, 100 bi-exp, 1exp+C and 1exp respectively. 50 Plots ‘b-d,’ show absolute intensities vs time, of 9 Intensity (a.u) Intensity 0 typical individual S-M traces of dGTP-Cy3/DNA 0 250 500 750 1000 (#2) and their sudden bleaching steps. d) Time (seconds)

Figure 35) Selected frames from dGtpCy3/DNA_sept movie showing decrease in fluorescent S-Ms due to photobleaching.

Figure 35): continued:

From left to right: Frame 1 showing Cy5-Primer/DNA locations under red laser. Frames 5, 65, 125, 250, 400, and 600 show dGTP-Cy3 positions either incorporated into the DNA-Primer or not. Again, not all the molecules were bleached but only those that did bleach were analyzed and their fluorescence profiles summed up to obtain the graph in Fig.34 (a). Scale bar shown is 5µm and sample area is 200 x 200 pixels or 20µm x 20µm.

Note: After observing the Cy5-Primer in frame 1 one can see a drastic drop in the number of fluorescing dGTP-Cy3/DNA molecules in frame 5. This corresponds to guanine’s superior quenching characteristics compared with the other nucleotides. After some time again one can see an increase in the number of fluorescent single molecules, frame 65-125, followed by the inevitable decrease in the number of fluorescent S- Ms due to photobleaching under continued illumination, frames 250-600.

There were 114 S-Ms selected in the above experiment out of about 275 correlated objects. 93 of these, or 82%, were initially quenched for some time before exhibiting sudden fluorescence and then being irreversibly photobleached after continuing illumination. Approximately half of the quenched molecules were non-fluorescent for up to 100 seconds with some being completely quenched for over 600 seconds. Quite a few of the fluorescent traces were not typically smooth and rectangular in profile but rather showed several distinct peaks which lasted either just 1 or 2 frames or perhaps as much as 20 to 30 frames. This might be due to what’s known as photoblinking of the fluorophore and is quite common among organic dyes. It is thought that the ‘on-off’ profile is due to the dye interchanging between singlet, (on) and triplet, (off) states. The longer lived triplet states requires the dye to undergo reverse intersystem crossing to get back to the singlet state again and this can be in the order of a few milliseconds to seconds depending on the molecular environment. For instance in the presence of oxygen the triplet lifetime can be reduced from 10’s of milliseconds to micoseconds as the molecular oxygen

80 quenches the triplet state faster [30]. Because of the rather long acquisition time of 2 seconds per frame it is thought that the majority of the dye molecules will experience the full range of dipole-dipole orientations under the laser illumination within 1 frame. Hopefully this would have the effect of screening out most of any biased orientations that would otherwise be unevenly excited under various polarization planes.

3.7 Preliminary results for incorporated dGTP-Cy5 with Oxygen Scavenger Solution

Figure 36) Graph showing bleaching lifetime of all Single-Molecules analyzed along with selected S-M traces of dGTP-Cy5/DNA in O2 scavenger.

Fluorescence intensity profile of dGTP-Cy5/DNA in O2 scavenger

0.6

data 0.5 Bi-exp+C 0.4 1exp+c 0.3 Bi-exp 0.2 1exp 0.1 Relative Intensity Intensity Relative 0 0 100 200 300 400 500 600 Time (seconds)

Graph showing the total ‘normalized to maximum fluorescence’ intensity vs time of all bleached single- molecules of dGTP-Cy5/DNA in Oxygen scavenger solution. Bi-and mono-exponential curves were fitted using a least squares fit analysis with chi-squared values of 0.82, 0.83, 0.83 and 0.83 for bi-exp+C, bi-exp, 1exp+C and 1exp respectively. Bleaching lifetimes were also relatively similar for each curve with values of approx. 220-250 seconds.

As can be seen from the above graph there is no apparent rise in fluorescence at all associated with dGTP-Cy5 in oxygen scavenger solution. The scavenger used was a glucose oxidase and catalase based solution to inhibit singlet oxygen formation or at least degrade it as soon as any was formed. Cy5 is known for somewhat erratic fluorescence and complicated bleaching behavior [30]. But as can be seen in this graph with scavenger present all 4 exponential type fits are almost identical in their chi-squared values. 137 molecules contributed to the graph out of 254 objects which corresponds to over 52% incorporation! Upon observing the movie there was no distinguishable increase the number of fluorescent molecules on screen. However, there was a considerable increase in the amount of what appeared to be ‘photoblinking’ i.e. rapid (within 1 or 2 frames) on off switching of fluorescent molecules. It is not clear if this is due to the scavenger solution or Cy5 itself.

It would appear that blocking singlet oxygen formation has a very significant effect on the type of fluorescence exhibited at the single-molecule level.

Figure 37 Selected frames from dGtpCy5/DNA O2 scavenger movie showing decrease in fluorescent S-Ms due to photobleaching.

From left to right: Frame 2 showing Cy3-Primer/DNA locations under green laser. Frames 7, 125, 250, 375, and 525 show dGTP-Cy5 positions either incorporated into the DNA-Primer or not. Again, not all the molecules were bleached but only those that did bleach were analyzed and their fluorescence profiles summed up to obtain the graph in Fig.36. Scale bar shown is 5µm and sample area is 150 x 150 pixels or 15µm x 15µm.

4 Discussion

4.1 Evidence for Guanine oxidation

As was mentioned previously in section 1.3 guanine is the most readily oxidizable base [35, 36] and therefore is expected to be most susceptible to damage by oxidation and radical forming processes. This indeed proves to be the case and there are many papers documenting the various guanine related adducts and lesions formed by these processes [10, 12, 20-23, 40, 49, 87-91]. Examples of the main observed guanine moieties from single-electron oxidation and singlet oxygen mediated damage along with hydroxyl radical damage are shown on the following page along with their various reaction pathways.

Again it should be noted that depending on whether or not the guanine base is in free solution or encompassed in DNA as a nucleotide, varying oxidative products are formed. This is understandable when comparing the nano-environment of a dGTP nucleotide wrapped up in a DNA helix with that of a free nucleotide. Obviously the free nucleotide is more exposed to singlet oxygen and other ROS in the solution compared to dGTP ‘shielded’ in the more hydrophobic interior of double stranded DNA. This has been shown experimentally by [92] with the guanine adduct 8’-oxo-dGuo in single stranded DNA being up to 7 times more sensitive to singlet oxygen than that in double stranded DNA. In double stranded DNA by far the main oxidized guanine derivative via singlet oxygen reactions is 8’-oxo-dGuo [89] and ¹O2 oxidation can lead to base damage and strand breaks which occur mainly at these guanine residue sites.

Figure 38) Diagram showing 11 of the main guanine adducts and oxidative lesions formed via one-electron oxidation, hydroxyl radical damage and ¹O2 mediated reactions in solution.

From the above diagram it is clear that guanine or dGuo (middle) can be prone to many different reactions within its surroundings. It is possible that a combination of these processes can attack guanine almost simultaneously. Diagram taken from [93] J. Cadet et al. /Mutation Research 531 (2003) 5–23

As can be seen from our DNA/Primer in figure 18) the DNA template (T404) is single stranded with the primer being double stranded. This means that the last incorporated nucleotide (dNTP-Cy3) onto the primer is somewhat more exposed to singlet oxygen attack compared to being in the middle of the primer sequence and perhaps less exposed if it were simply on a single strand.

Singlet oxygen is known to predominantly attack guanine residues in DNA in equal amounts [41, 92-94] but is not necessarily excluded from damaging the other bases. This could explain the far fewer cases (app.5%) of delayed fluorescence and partial quenching of the other dNTP-Cy3 analogs that would be almost impossible to detect at ensemble level experiments. However with regards to one-electron oxidation processes there is some selectivity as to what sites are more prone to oxidation with 8’-oxo-dGuo sites being the most readily oxidized even compared to nearby 5’-GG-3’ and 5’-GGG-3’ sites [92].

From looking at figure 36) and comparing the fluorescence profile of dGTP-Cy5 in oxygen scavenger solution with that of the dGTP-Cy3 graphs it is clear that there is no increase in fluorescence intensity linked to dGTP-Cy5 in scavenger. Whilst it must be noted that Cy5 and Cy3 are not exactly the same they are of course very closely related in their chemical structure (see figure 17) and might be expected to behave alike under similar experimental conditions all be it with Cy5’s much faster bleaching lifetime. Therefore it would appear that the lack of singlet oxygen available in the surrounding medium, due to the scavenger solution, seems to prevent the complete initial quenching effect of dGTP on the dye’s fluorescence. Obviously this experiment needs to be done with dGTP-Cy3 in scavenger solution to observe the effects of removing singlet oxygen formation from the environment.

Previously Douki and Cadet ’99 [32] did an experiment showing how the bases can be modified by photo-induced oxidative processes, namely TYPE I (via radical/single

86 electron transfer reaction) or TYPE II (singlet oxygen,1O2, mediated reaction) PHOTOSENSITIZATION. They attributed the majority of damage to the guanine base from all 3 photosensitizers used due to its lowest oxidation potential and charge transfer properties within the DNA helix, predominantly proton coupled transfer due to hydrogen bonding between complementary bases [18] and radical migration through the π -stack [46, 50, 86]. It is therefore this combination of type I and II photo-induced oxidative processes that I believe are responsible for the differences observed between the fluorescent profiles of dGTP-Cy3 in DNA compared with the other dNTP-DNA configurations.

Figure 39) Diagram depicting possible reaction pathways explaining observed S-M fluorescence profiles of Cy3-dGTP.

fast -· +· slow Cy3*dGTP Cy3 dGTP Cy3* (dGTP)Oxy

(1) Complete (2) Quenching Fluorescence

fast Cy3-dGTP Cy3-(dGTP)Oxy

Single-Molecule Traces dGtpCy3_sept

) 300 (3) Dye

a.u 250

(

Photobleached

y 200 (2) 150 100 (1) (3)

Intensit Cy3Oxy (dGTP)Oxy 0 0 250 500 750 1000 Time (seconds)

Stage (1) represents complete quenching of fluorescence by the much faster process of electron transfer (see also figure 40). Upon irreversible oxidation by singlet oxygen quenching by electron transfer is stopped, stage (2). After further singlet oxygen build up the dye is then irreversibly photobleached and fluorescence ceases.

Figure 40) Diagram showing in more detail the electron transfer process between an acceptor and donor molecule.

The above diagram represents stage (1) in figure 39). The Cy3 dye acts as acceptor and the dGTP nucleotide as electron donor. Both ground and excited state energy levels are shown for both as well as singlet spin states of electrons involved. Due to charge recombination the process is cyclic and hence fluorescence is not permitted as electron transfer occurs on a much faster time scale (pico-seconds compared to nano-seconds).

In this case the Cy3 dye is believed to act as an electron acceptor and the dGTP as donor. Upon illumination the Cy3 electron is excited and another electron from dGTP is transferred to the now empty ground state level of Cy3. This prevents the excited electron from decaying back down to the ground state thereby preventing fluorescence (total quenching as seen in (1) in previous figure. The reason for total quenching becomes clear when one takes into account the relative reaction rates for electron transfer, 10¹¹/sec, compared to fluorescence, 108 /sec. The excited electron just doesn’t have enough time to get rid of its energy by photon emission as fluorescence.

After some time the electron transfer process stops and fluorescence suddenly appears. This can be explained by the slower more gradual build up of reactive singlet oxygen which chemically alters the guanine moiety in such a way that it can no longer act as an electron donor to the Cy3 dye. In other words it is irreversibly oxidized by the ¹O2.

This is equivalent to part (2) of figure (39). Now the dye continues to fluoresce until eventually it too has been irreversibly photobleached due to further build up of singlet oxygen.

The reaction kinetics of the previous two diagrams can be explained below:

4 4 K1=10 11 -2 K4=10 K2=10 K3=10 Cy3* G(Oxy) Cy3 G Cy3* G Cy3-· G+· Cy3 G(Oxy)

8 K-1=10

11 Cy3(Oxy) K-2=10 G(Oxy) where the rate equations are given as: • =+∗ −• +• − [Cy3 G]kCyG−−12 [ 3 ] kCyG [ 3 ] kCyG 1 [ 3 ] [1] • ∗∗∗=− − [Cy3 G]kCyG12 [ 3 ] k [Cy3 G] k− 1 [Cy3 G] [2] • ∗ -++•+ =− −•• − −•• [Cy3 G ]kk22 [Cy3 G]− [Cy3 G ] k 3 [Cy3 G ] [3] • = -+•• ∗ [Cy3 G(Oxy)]k3 [Cy3 G ]+ kk−44[Cy3 G(Oxy)]- [Cy3 G(Oxy)] [4] • ∗∗∗= [Cy3 G(Oxy)]kk44 [Cy3 G(Oxy)]-− [Cy3 G(Oxy)]- k 5 [Cy3 G(Oxy)] [5] • = ∗ [Cy3(Oxy) G(Oxy)]k5 [Cy3 G(Oxy)] [6]

The above equations can be grouped together and simplified as follows: • =− [Cy3 G]kCyG1 [ 3 ] [7] • = [Cy3 G(Oxy)]kk12 [Cy3 G]- [Cy3 G(Oxy)] [8] • = [Cy3(Oxy) G(Oxy)]k2 [Cy3 G(Oxy)] [9]

Equation [7] describes the exponential decrease in the amount of original Cy3-dGTP present and equation [9] represents the increase in irreversibly photobleached Cy3-dGTP product i.e. Cy3(Oxy)dGTP(Oxy). The amount of bleached product increases until all the original molecules analyzed in the graphs are irreversibly destroyed. Equation [8] is a combination of [7] and [9] and best describes the rate at which oxidized Cy3-dGTP is formed. The solution to eqn. [8] is a double exponential which is why the observed data curves for the dGTP experiments are best fitted to the double exponential equation (see figures 32a and 34a).

There are of course other possibilities of what can happen due to such a wide variety of oxidative processes and guanine derivatives being allowed, as can be seen in figure (38). For example a proton coupled transfer process to the complementary cytosine can occur leaving a neutral guanine radical or perhaps the excited electron in Cy3 can, through charge transfer, return to the now radical cation dGTP donor ready to repeat the process. Because singlet oxygen is known to be formed from triplet-triplet reactions with many dyes and molecular ground state oxygen [30, 63] as well as thermal excitation it is more than capable of being a major oxidant at the single molecule level. In fact it only takes 0.98eV to excite a triplet state oxygen molecule to its excited singlet state and this is quite possible when colliding with a dye molecule in its triplet state. The triplet dye then returns to the singlet ground state and can be excited once more. Thus another source of singlet oxygen formation can be produced locally within the dGTP’s nano- environment. Molecular oxygen greatly reduces the triplet lifetime of the dye from milli- to micro-seconds thus further enhancing the rate of singlet oxygen formation. As already explained a major effect of singlet oxygen reactions on guanine in DNA is the formation of the lesion 8’-oxo-dGuo which actually has a much reduced (between 0.58V and 0.75V v NHE less) oxidation potential than that of guanine in DNA (1.29V v NHE) [35]. This lesion is even more susceptible to further one electron oxidation and singlet oxygen reactions which are becoming more of an interest in recent times.

It is also possible that the 8’-oxo-dGuo that has been created from type I and type II photo-induced oxidative processes [16, 31, 87] can donate electrons to the dye’s ground state as it has a much reduced oxidation potential even compared to guanine. Upon repeated cyclic electron transfer processes and the build up of singlet oxygen from dye- molecular O2 reactions the total quenching ability of dGTP or 8’-oxo-dGuo molecule stops. This is most likely due to an irreversible oxidative guanine derivative formed such as oxazolone which is an end product created when reacting with singlet oxygen, oxaluric acid is also formed [89, 95, 96]. Once the now irreversibly oxidized guanine lesion can no longer quench the dye’s fluorescence there opens up the possible pathway for a photon to be emitted from the Cy3’s excited singlet state allowing the excited electron to return back to its ground state and fluorescence to occur. This can explain the sudden appearance of fluorescence in the dGTP-Cy3/DNA experiments. Of course singlet oxygen can still be formed from triplet-triplet dye interactions and its continuing build up along with a chemical reaction between the triplet-dye-molecular oxygen complex will eventually damage the Cy3 molecule leaving it irreversibly photobleached. The exact mechanisms behind the reaction pathways for photobleaching are being better understood but still lack the analytical power to cover all dyes [30, 65, 97, 98].

It is not clear exactly what the ground and excited state potentials of the Cy3-dGTP molecules actually are but they must be thermodynamically and electronically compatible in order for the electron transfer processes to be allowed. It is interesting to note that at the time of writing it is no longer possible to buy dGTP-Cy3 from PerkinElmer Life Sciences. It is conceivable that the seemingly irregular fluorescence properties were deemed unreliable compared to the other non-quenchable dNTP-Cy3 analogs.

4.2 New light on interpretation of photobleaching kinetics in single- molecule fluorescent traces.

Recently there has been evidence of what are known as long lived dark states [97, 99] that are linked to more stable radical anion formation of the dye, in their case, Rhodamine 6G. They imply that these other metastable states can greatly alter the photobleaching profiles. Although Zondervan et al do not specifically address Cy3 it is interesting to note that both they and Kohn et al [30] show the case for non-exponential bleaching dynamics and highlight other metastable states that are intermediates in the bleaching kinetics of single-molecules; radical anions in the case of Zondervan and ‘twisted’ isomers of Cy3 in Kohn’s case.

Even more recently Hoogenboom et al [100] states that these long lived radicalized dark states are the main precursor to photobleaching, not necessarily the triplet states and that they obey power-law [98] statistics. They attribute the electron tunneling from the surrounding medium to the excited state (either singlet or triplet) of the dye as the predominant pathway to photobleaching of the molecule.

It could entirely be possible that a radical anion Cy3 dye state can also be formed by single electron transfer from an electron donor such as a nucleotide (dGTP or 8’-oxo- dGuo). This electron, after transfer to the dye can then react with the water or oxygen in the surrounding medium and release more ROS or perhaps transfer back to the ground state of the nucleotide again.

What is clear is that interpretation of single-molecule data is not as simple as might previously have appeared and recent experimental observations are shedding new light on the statistics behind the single-molecule experiment.

5 Conclusions and suggestions for future work

I have shown that there is a marked difference in the fluorescence properties of dGTP-Cy3/DNA compared to the other nucleotide configurations at the single-molecule level. By observing statistically appropriate numbers of S-M traces it is clear that dGTP- Cy3 exhibits extreme and often total quenching of fluorescence for various times under laser illumination. After some time this complete quenching stops and normal fluorescence abruptly appears followed by eventual photobleaching of the Cy3 dye upon continued illumination. We believe this delayed fluorescence due to total quenching is related to guanine’s much lower oxidation potential compared with the other nucleotides in DNA. It is this low oxidation potential that allows it to be prone to excessive oxidative damage by various processes such as one-electron oxidation via radical formation and singlet oxygen mediated damage. These processes are simulated in our experiment through a photo-induced oxidative method using Cy3 as the possible electron acceptor as well as being a singlet oxygen producer via triplet state interactions with the surrounding molecular oxygen. These are known as type I and type II photo-sensitization processes. It is the combination of these 2 pathways that can lead to irreversible oxidative damage of the guanine base in the form of an end product such as oxazolone. Once this occurs it can no longer quench the fluorescence by electron transfer or react further with singlet oxygen and this allows the sudden onset of fluorescence as evidenced in the dGTP-Cy3 single-molecule traces. There is still the possibility of misinterpretation of the single- molecule data as some other unseen factor perhaps due to the dye itself. However, the fact that dGTP-Cy3 is so much different compared to the others lends weight to the fluorescence variations being due to the nucleotide rather than the dye.

There are a few experiments that can be done to help elucidate the findings within this dGTP-Cy3/DNA experiment. One obvious experiment is to repeat the Oxygen scavenger experiment but with Cy3 instead of Cy5. This would have been done if it were not for the weakened intensity of the green laser. The time taken to bleach enough S-Ms

93 was therefore much too long and was beyond the capacity of the correlation software. With a higher intensity laser it will be possible to see how the prevention of singlet oxygen affects the S-M fluorescent traces.

Another experiment to ascertain the affects of singlet oxygen would be to incorporate several non-labelled nucleotides after the fluorescent dGTP. This would shield the nucleotide more from the surrounding medium as it is deeper in the double stranded primer sequence. By again analyzing the amount of quenching it might be possible to compare the effects of singlet oxygen damage to various exposed and more protected dGTP configurations.

If an end by-product like oxaluric acid is given off then it should be possible to detect it in the sample solution. Due to the very low concentrations however it might prove difficult to detect these amounts. On the other hand there is no reason not to repeat the experiment at higher concentrations having already observed the fluorescent profiles at the S-M level and bleach the sample (using a higher intensity illumination). This should increase the amount of end product formed and facilitate its detection.

One last more complicated experiment would be to make use of certain DNA glycosylase repair enzymes that cleave off oxidized guanine nucleotides at the deoxyribose sugar ring. By flushing in new dGTP-Cy3 or perhaps dGuoCy3 (if possible!) with DNA polymerase and comparing before and after fluorescence movies using the correlation software it should be possible to ascertain if the original dGTP was oxidized and removed. Any new incorporated dGTP-Cy3 molecules would again exhibit the delayed fluorescence as previously shown. Obviously great care must be taken in flushing in and out the reagents without disturbing the position of the sample more than a few pixels in length.

In short there is much documented proof for guanine oxidation within DNA and its relevant importance in understanding the mechanisms that lead to base lesions and mutations which are thought to be major underlying precursors in aging and chronic diseases such as cancer. It is quite possible that we have indeed observed single DNA, guanine specific, irreversible oxidation events using single-molecule fluorescent measurements.

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