Development of a microfluidic platform for integrated DNA sequencing protocols

Development of a Microfluidic Platform for Integrating

Nanoliter DNA Sequencing Protocols

By

Mayank Kumar

B.Tech. Department of Mechanical Engineering (2004) Indian Institute of Technology Kanpur, INDIA

Submitted to the Department of Mechanical Engineering in Partial Fulfillment of the Requirement for the Degree of

Master of Science in Mechanical Engineering at the

Massachusetts Institute of Technology February 2007

© 2007 Massachusetts Institute of Technology

Signature of Author

Department -Meeanical Engineering January 19, 2007

Certified by

Todd Thorsen Assistant Professor of Mechanical Engineering Thesis Supervisor

Accepted by

Lallit Anand Chairman, Department Committee on Graduate Students MASSACHUSETS INSTTUTE

MIT

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Development of a Microfluidic Platform for Integrated DNA

Sequencing Protocols.

by

Mayank Kumar

Submitted to the Department of Mechanical Engineering on January 19, 2007 in Partial Fulfillment of the Requirements for the

Degree of Master of Science in Mechanical Engineering

Abstract

This thesis describes the design and development of a microfluidic platform to reduce costs and improve the quality of in the DNA sequencing methodology currently implemented at the Broad Institute in Cambridge, Massachusetts.

The Sequencing Center at the Broad Institute currently generates an average of 130 million bases per day with an average read length of 800. This is enabled by the successful preparation and detection of over 97,000 unique samples. Most of the cost per sample is tied up in expensive proprietary reagents utilized in the various reactions comprising the preparation process. Through the application of microfluidics, the possibility of drastically scaling down the amount of proprietary reagents is explored.

Stamp-sized elastomeric polydimethylsiloxane (PDMS) microfluidic devices were developed and microfluidic sample manipulation techniques were standardized. Using these devices and techniques, an attempt was made to adapt the various components of the sequencing process to the microfluidic platform.

Work within the scope of this thesis is focused on the adaptation of the commercial sequencing protocols, which are labor intensive, consume costly reagents and serve as limitations for high-throughput parallelization of the process. The first is the amplification reaction. By scaling down the process from a plate-based format to an integrated microfluidic device, amplification reagent consumption was reduced by two orders of magnitude while maintaining the quality and length of the sequencing reads (with the subsequent sequencing reaction run off chip). As a follow up project, an attempt was made to scale down the Sequencing Reaction, which, in spite of limitations, suggested a good path toward the eventual development of an integrated microfluidic device for the preparation of running the complete sequencing reaction protocol on-chip.

Thesis Supervisor: Todd Thorsen

Acknowledgements

First and foremost, I thank Professor Todd Thorsen for reposing faith in me and entrusting me with the responsibility of working on this project. I am very grateful for being able to use the resources of his lab at MIT. He was encouraging at my successes and equally understanding at my failures. None of the work here in this thesis would have been possible without his extremely friendly and affable nature. I feel very fortunate to have had his guidance and creative suggestions at so many critical junctures.

I thank the Broad Institute for providing me with the most state-of-the-art resources in

Bioengineering research. It was a great privilege to have worked on the most advanced instrumentation and methods. The most valuable, however, was the help and advise of a great team of researchers in the Production Sequencing Development section of the Broad Institute where I was employed. I have no words to express my gratitude to Joe Graham who was my research supervisor at the Broad Institute. None of this would have been possible without the calming assurance provided by his insightful suggestions, care and understanding. I heartily feel that his affectionate behavior was always like an elder brother than a supervisor.

In addition, I thank Andrew Barry, Sheila Fisher and Susan Faro at the Broad Institute. Andrew is the most cheerful guy I have ever met. A shining beam of enthusiasm, he was ever ready to help me regardless of the complication of the task at hand. I am grateful to Sheila for providing me with her invaluable advise in the myriads of times of crisis and for helping me with the bioengineering techniques which were completely foreign to me to begin with. Susan's readiness to help coupled with her motherly care and affection provided me with the most affable work environment possible. I will also like to thank my labmates at the Thorsen Lab. I am grateful to J.P. Urbansky for helping me out with the nitty-gritties of microfluidics on numerous occasions. His prior experience in the issues I encountered made my work so much easier.

I will also like to thank Leslie Regan, Joan Kravit and the entire graduate office for

helping me out since the very beginning of my stay at MIT. They made MIT feel like a second home to me. For the same reason, I thank my friends Siddarth, Piyush, Kaustuv, Ajit and many others who made me feel homely and cared for.

I thank my family for always providing great strength to me in my hardest times. No

words come close to the amount of love I have received from my parents since times of which I don't even have a conscious remembrance. The love from my mother, father and sister is the most valuable thing of my life and whenever I think of that, the day-to-day problems and challenges suddenly become very small and easily manageable.

Last but not the least, I thank that Source, the Ultimate Reality from and in which my personality and its talents and capacities have sprung. I know that my abilities are mine

by accident and not by choice. I am grateful for this life and its enriching experiences.

Contents

1. Introduction ... 9

2. D evelopm ent of Basic M icrofluidics ... 23

3. Am plification ... 37

4. Sequencing Reaction ... 63

5. Introduction ... 77

APPENDICES 1. Fabrication ... 81

2. LABVIEW Controls and Instrumentation ... 85

Chapter 1

Introduction

The purpose of this thesis is to propose cost reduction and quality improvement techniques in the DNA sequencing methodologies at the Broad Institute through the adoption of a microfluidic platform.

The DNA Sequence

To set the goals and inspiration of this work, we need to first introduce the concept of the DNA sequence. The deoxyribonucleic acid (DNA) is a double-stranded molecular chain in the nuclei of our cells that carries the genetic information that is subsequently translated into proteins that not only engage in complex regulatory pathways at the cellular level, but form higher order structures like tissue and organs. Its shape is a double helix (Figure 1.1), consisting of a string of simple units called nucleotides, which are held together by a backbone made of sugars and phosphate groups.

The backbone consists of four types of molecules called bases and it is the sequence of

these bases that encodes information. The four bases as shown in Figure 1.1 in different

colors are Adenine (A), Cytosine (C), Guanine (G) and Thymine (T).

LI

K

I

~

Figure 1.1 DNA Structure details

Hence, the DNA sequence is a sequence of these letters A, T, G and C, which are

representative of the bases in the DNA chain. DNA sequencing is the process of figuring

out the sequence of these bases through chemical reactions and the use of appropriate

technology.

is six orders of magnitude larger than this read length. This is the mitigating circumstance, which has led to the current technique employed for large genome sequencing. This technique consists of shattering the genome of interest (with or without some level of preliminary sorting) into small cloneable segments, sequencing the ends of these segments, and utilizing an assembly algorithm coupled with a large amount of processing power to reassemble the genome based on overlaps in this sequence.

The Sequencing Center of the Broad Institute currently generates an average of 130 million bases of sequence every day. This is enabled by the successful preparation and detection of over 97,000 unique samples. Each of these preparations carries an associated cost which is the sum of all reagents, consumables, labor, and detection time. The value of a high throughput facility is in the ability to quickly amortize fixed costs for start-up such as automation, process tracking software, initial R&D, etc.

When addressing process improvement, an obvious target is the reduction of consumable costs (assuming no increase in detection capacity). For the sequencing process, most of the cost per sample is tied up in expensive proprietary reagents utilized in the various reactions comprising the preparation process. The most straightforward way to reduce the costs associated with these reagents is to reduce the amount of reagent used per reaction or eliminating steps all together.

The purpose of this project is to propose cost reduction and quality improvement by directly cutting down the reaction volumes by at least two orders of magnitude through the adoption of nanoliter sample preparation platforms.

At the Broad Institute currently, the limit on sequencing is detection. The production process in place can easily, in a 40-hour week, generate enough samples to keep detection machines running 24 hours a day, 7 days a week. In such a scenario, it does not make much sense to concentrate efforts on decreasing process time for sample preparation. However, introduction of microfluidics will easily serve to reduce the time and complexity of the process as well.

Currently, the sequential series of steps involved in sample preparation, amplification and sequencing reaction take place in discrete robotic stations. In addition, there is the tedious manual transfer of reaction plates from one station to another. These are certainly avoidable issues in labor efficiency. With the development of a suitable microfluidic platform, electronic automated controls can make it possible for the above mentioned series of steps to be performed in a single device sequentially without the need for manual interference. Hence, apart from reduction in reagent consumption, microfluidics is extremely useful in simplifying the entire macroscopic scheme of operations.

The quest to miniaturize lab processes has been ongoing in the biotechnology industry for years, reflected in the evolution from 96 to 384 and then to 1536 well plates. When operating with reactions at the pL level, environmental factors begin to play a larger role and the need to control evaporation, condensation, and contamination becomes a necessity. In order to overcome with this issue, scientists are relying more and more on microfluidic technology. The utilization of this type of technology is already pervasive in the industry and can be seen in such applications as HPLC, Micro Electro Mechanical Systems (MEMS), Lab-on-a-Chip, and Capillary Electrophoresis (CE). A strong

argument can be made, in fact, that it was the advent of CE for DNA sequencing that enabled the Human Genome Project to be finished years ahead of schedule at a reduced cost due to the increased ease and throughput of this technology.

Numerous research groups have worked significantly towards miniaturizing the process of DNA sequencing. Much work has been done on taking the electrophoretic separation and detection part of the sequencing process to the microfluidic realm, for example by Mathies group. Also, we have seen great progress in making the DNA amplification technology of Polymerase Chain Reaction (PCR) possible on nanoliter and even picoliter scale. This is significant since PCR is still used at some places as a part of the sample preparation protocols. But we are yet to witness systematic efforts to sequentially map

For example, at the Broad Institute, the sample preparation roughly consists of bacterial cell culture, followed by Rolling Circle Amplification (RCA), Sequencing Reaction and purification. Similar processes are in place at other major sequencing centers including the Joint Genome Institute (JGI) in California. Still we have not seen any cases of RCA being implemented at nanoliter scale to extend the possibility of its integration with the other sample preparation processes at a microfluidic scale. Of course, complete miniaturization of the process demands answering many issues including modifying the dispensing mechanisms, bacterial transformation and library construction methods and also probably the detection mechanism.

In this thesis, the efforts made at bringing down RCA and subsequently the Sequencing Reaction to the nanoliter scale will be outlined. Chapter II discusses the development of fundamental microfluidic fabrication and functional aspects, which will be required for the prospective nanoliter sample preparation platform. Chapter III includes extensive experimentation on the DNA amplification reaction on the nanoscale leading to a set of optimized conditions and protocols on reagent consumption and template preparation. Chapter IV similarly discusses the efforts made on making a nanoliter Sequencing Reaction on a PDMS microfluidic format possible. But first we discuss here the protocols already in use in the DNA Sequencing line at the Broad institute.

Library Construction

The process by which these DNA samples are prepared for sequencing begins with the construction of a library. This is schematically shown in Figure 1.2. The first step is shattering the genome of interest into short fragments, generally about 1000 base pairs in length, using restriction enzymes shown as yellow beads in Figure 1.2. Then the same enzyme is used to cleave the plasmid of interest at a similar site. The plasmid is a circular piece of double stranded DNA generally about 2000 to 5000 base pairs long. The small

The remarkable property of plasmids is that the bacterial cells do not differentiate them

as foreign for purposes of replication. Moreover, the plasmids multiply within the host

cells in the absence of cellular multiplication. The vector DNA is then inserted into

bacterial cells (transformation). The bacterial cells are then spread onto agar plates

where, provided they were successful transformed with an inserted vector, they propagate

into colonies of thousands of identical cells, each containing an exact copy of the inserted

DNA.

These colonies are then detected through standard imaging techniques and

transferred into sample plates via automated picking machines for purification and

sequencing. This is a labor intensive and time-consuming process, but it is necessary to

separate and amplify the DNA being interrogated into unique clones for sequencing.

EIZ~

Chromosomal DNA

Plasmid

Insert Target DNA

Host Cell

Transformation

Plasmids Multiply

Host Cells Multiply

Colonies Grow

Figure 1.2 Recombinant DNA Technology

Once isolated, the bacteria are subjected to a series of purification steps to isolate and

amplify the inserted DNA of interest followed by a Sanger sequencing reaction to

generate detectable product. The amplification takes using TempliPhi 10000 (Amersham

Biosciences), which makes use of the Rolling Circle Amplification technique discussed

in detail in Chapter 1II. Currently, the whole process is accomplished as illustrated in

Figure 1.3, with samples migrating through a heavily automated process in 384 well

plates and undergoing pL volume reactions.

Inoctd at Ion DenaT ure Add TPh i A mplify/ bIut in

Heat Kill

Transfer &lycerol Denature at Add Tphi Amplify at D Iuto with

and Add Denaturing 95C for from Ch iled 3 I 6hr 3rUL wator

Buffer 5min. Resevoir 6*C/]yn

S eq. Set -Up The rmocycling ETOH Precp. EDTA Elut ion bete Cion

&4WE~

4W

4Wli 4W

W

4W

4W

4W

4Wk

TrnfeWo e 9 5c5 'C- Add Ethanol Load on ABI

Transfer to Se . Plates 60"C 40 and Add EDTA 3730

& Add Mi oydes Centrifuge Detector

Figure 1.3 Complete Sequencing Process at the Broad Institute

The current process steps are:

1.

.5ptL of the isolated bacterial culture from one well of a 384 well glycerol plate

(200pL max. volume) is combined with 2pL of denaturing buffer in a new 384

well conical bottom plate (35 ptL max. volume).

2. The plate is then sealed and run through a conveyor oven which raises the sample

temperature to 90'C for 3 minutes.

4. The plates are then resealed and placed in a chilling incubator which heats the samples to 30'C for 16 hours then raises the temperature to 65'C for 1 minute before cooling the samples to 10'C, which they remain at until removed and placed in a refrigerator to await sequence set-up.

5. The plates are then diluted in a two-step process the first step of which adds 14.5RL of water, mixes, aspirates 15RL and deposits that to waste. The second step of the dilution adds another 15pL of water to the remaining 5pL of dilute sample and mixes.

6. 1.3pL of the dilute sample then is transferred into two wells of a new 384 conical

bottom plate (35ptL max. volume). .75ptL sequencing mix consisting of Big DyeC reaction mix and primers specific to either the 'forward' or 'reverse' site of the vector then gets added to each of these wells.

7. These new plates are then sealed and placed on thermocyclers where they undergo 25 cycles of 96'C for 20 seconds to 50'C for 15 seconds to 60"C for 4 minutes.

Once the cycling is complete the plates are stored in a refrigerator.

8. The seals are removed and 14 tL of Ethanol is added to each well. The plates are

resealed and centrifuged at 2000 rpms for 2 minutes. The ethanol is then removed

by inverting and tapping the plates. The plates are then stored in a warm room.

9. The plates have 10pL of EDTA added to each well, are spun, sealed, and loaded

DNA Amplification

Prior to the Sequencing Reaction, it is necessary to amplify the DNA of interest. This

amplification serves two purposes:

1. When isolating the DNA from bacterial cells, selectively amplifying the DNA

allows it to become the overwhelming component of the reaction mixture thereby

eliminating the need to purify out any bacterial genomic DNA, proteins, or other

agents that may interfere with the sequencing reaction later on.

2. Amplification of the DNA of interest allows for a uniform amount of input into

the sequencing reaction. The amplification step is achieved through the use of

TempliPhiC which utilizes the Phi29 enzyme for rolling circle amplification

(Figure 7), explained in much greater detail in Chapter III.

Figure 1.4 Rolling Circle Amplification

One thing to take note of with regards to the amplification reaction is the dilution step.

This is essentially a 1: 615 dilution of the amplified product prior to input into the

sequencing reaction. Simply reducing the volumes associated with this reaction should

make it possible to eliminate this dilution step all together, a feasible adaptation for

microfluidic devices.

Prior reagent minimization efforts have been conducted by S. Fisher et al. [1], which

reduce the volume of TempliPhi reagent used down to 100nL while keeping the overall

Figure 1.5, acceptable quality sequence can be obtained from template generated from

only 400 nL of TempliPhi without making any other adjustments. This shows the large

dynamic range of the reaction and suggests that further optimization is achievable by

lowering the overall reaction volume. It is interesting to see how this volume actually

scales down under real microfluidic experimentation in Chapter III.

50 45 40 35 30 20 15 10 5 0 0 CP (0 0 a) 0a C6 CC) a) - a) Volume TPhi (nL) CO 0 1903 LO CO CD

0-Figure 1.5 Tphi Volume Reduction

Sequencing Reaction

the process using the minimum input material required by the ABI 3730xl sequencer to

determine the extent to which the process can be scaled down.

It has been shown by J. Meldrim[2] that the current process yields sequencing product in

excess of what is required for detection. The data generated shows that a significant

quantity of acceptable quality sequence (Phred 20 standard) can be obtained using the

product generated by the current 2pL sequencing reaction dilute down to only 2% of its

original concentration (Figure 1.6).

Theoretically, this would mean that a 40nL

sequencing reaction would yield sufficient product for detection. This possibility is

systematically explored in Chapter IV.

800

700-

I

400-

S300-

6

Dilution

Figure 1.6 Phred 20 for dilution series performed on a single sample

This analysis is not that straight forward; however, for one thing, the data also showed a

significant decline in the detection intensity for the dilute samples (Figure 1.6). This

means the reliability of generating consistent reads would be challenged by the optical

sensitivity of the detectors. This may be due to the sample loading interface to the

detectors, which currently requires a minimum elution volume for electrokinetic

injection. Effectively injecting the largest amount of sample possible, yielding the

greatest intensity, would most likely require a modification to the injection interface to

reduce the necessary injection volume. Also, reaction kinetics cannot be overlooked

when dealing with reactions at this volume.

The Dispensing Problem

The scale down as shown in Figures 1.5, 1.6 is possible only with specialized dispensing

devices. However, with the current dispensing machines in use at the operating stations at

the Broad Institute, the Equator and Multimek, it is not possible to go down to the

nanoliter scale and utilize the complete dynamic range of the reactions. Hence, even after

knowing that the different reactions involved in DNA sequencing can be scaled down to

the nanoliter scale, it is not possible with the existing dispensing instrumentation to

utilize this potential advantage. A methodology for nanoliter dispensing is required

which, as described in Chapter II, can be easily afforded in a microfluidic device.

The Possibilities

otal Volume 2.5

*Based on 1/64th BigDye concentration

)ilution /olume uL 0.5 ).5 2 3 74.5 80 0.63% 2.50% 3.75% 93.13%

Table 1.1 shows an initial estimate of the possibilities afforded by switching to a

microfluidic platform. These calculations are basically linear and are roughly based on an

elementary study of how the reads scale down when the amount of sequencing product is

reduced in a 384 well plate reaction. Beginning with the proposed sequencing reaction

volume of 40nL (2% of the current volume) and extrapolating out the required volumes

for the rest of the process keeping the ratios the same and eliminating the dilution, the

new 'microfluidic' process would consist of the volumes reflected in Table 1.1.

If

successful this would equate to a 98% reduction in Big Dye volume and a 99.94%

reduction in TempliPhi volume.

Picking Colonies Denaturation TempliPhi Addition

Amplification and Heat Kill Big Dye Addition Thermocycling Figure 1.7 Operating stations for various processes with robotic automation

Hence, resorting to a microfluidic platform has great potential to cut down reagent costs.

Furthermore, the whole sequence of operation can be greatly simplified with great

advantages in labor efficiency. As shown in Figure 1.7, the current process takes place

sequentially at several robotic stations requiring manual labor at several places. With

microfluidics it is possible to being all these steps potentially to a single point of

operation. This will greatly simplify process controls.

References

[1] Fisher, S., et al., Experiment conducted by the Technology & Development group of

the Broad Institute, 2003.

[2] Meldrim, J., Limit of Detection, Technology & Development at the Broad Institute

Presentation, 2004

Chapter 2

Development of Basic Microfluidics

Preliminary Device Design

The PDMS microfluidic devices were fabricated through the technique of soft lithography. [Appendix I]. In this chapter, we focus on the design of microfluidic circuitry in terms of the geometry of the flow channels and also on the protocols for some basic operations required to manipulate fluidic reagents on the nanoliter scale. The layout of devices is designed in Adobe Illustrator.

These devices are dual-layer devices having a flow layer for fluid flow and a control layer incorporating valves for controlling the flow. The fabrication and control using Labview of these devices is explained in Appendices I and II.

The first design as shown in Figure 2.1 was an ambitious one. This design aims to accomplish all the three prospective stages of sample preparation including amplification, sequencing reaction and purification within a single reaction chamber, which happens to

be the peristaltic rotary mixer. Moreover, it was proposed that the accurate metering of all

the reagents involved would also be accomplished in the same design.

-+ Cross Injection for metering definite amount of Template, Templi-phi and Big Dye, Primer.

-+ Dilution water forces these fixed amount of reagents in the rotary mixer. -- i12 Valves Template Force Template +* Templi-phi First Dilution Templi-phi

4F

Big Dye, Primer

> Total Sample Volume at Prior to injection = 50 nI.

--> Template : 0.4 ni, Templi-phi: 2.5 nI

Big Dye: 19 nl , Water: 28 ni

Beads + Alcohol Waste out Amplified Template -- Peristaltic Pump

I

1

Figure 2.1 makes the proposition quite clear. To the left side of the design, we would have the template (0.4 nl) and templi-phi (2.5 nl) accurately metered. They would then be forced by dilution water into the bottom left portion of the rotary mixer as shown in Figure 2.1. The reactants would be securely 'valved-off from both sides and would occupy merely 1/4 h of the 50 nI reaction chamber.

The mixing takes place by diffusion. Taking the diffusivity of average biomolecules in water as parameter, it was found that 7 minutes would be taken for complete diffusion mixing between the template and the T-phi. Considering that the amplification reaction itself consumes 16 hours of time in the macro-scale reaction, the diffusion time scale is really negligible to the reaction time scale.

After the amplification reaction is over, the requisite amount of big dye (19 nl) is metered in the bottom portion on the design (Figure 2.1) and forced in into the remaining 3/4th

portion of the rotary mixer by dilution water (18.5 nl). The volumes of reactants can be controlled using the cross-injection metering scheme. Figure 2. 3 illustrates the metering process in detail. The accuracy of metering really depends on the accuracy of the whole soft lithography process, which is discussed later in this thesis.

The two closed valves on both sides now only separate the adjacent volumes of amplified template and diluted Big Dye. Both these valves also form two of the three valves constituting the peristaltic pump. Three valves occurring in series on a channel when activated in a suitable pattern constitute a pump. Now, pumping is turned on by the mechanism as outlined by Unger et. al.[l]. Peristalsis is typically actuated by the pattern

101, 100, 110, 010, 011, 001, where 0 and 1 indicate "valve open" and "valve closed,"

respectively. The Labview control required for this operation is systematically discussed in Appendix 2.

The mixing would take quite bit of time here. The radius of the circular mixer here is 4 mm and peristaltic pumping at the operating valve pressure of 10-15 psi is barely

sufficient to support mixing in such a large volume. This deficiency is improved upon in the later designs.

The microfluidic device is subsequently placed on a flat-plate thermocycler for the sequencing reaction to take place. After these proposed sequence of steps, the DNA sample still remains to undergo purification in order to remove the excess ddNTPs from the Big Dye mix. Another inlet purges out 1/5th of the sequence reaction products, as shown in Figure 2.1, and replaces with magnetic DNA affinity beads. The circular chamber is again sealed and mixing done.

Subsequently, the DNA is sticking on to the beads. The chip is then placed onto a magnetic 96 well plate counterpart leading to the beads being immobilized in-situ. Water is flushed through the circular chamber cleaning away the impurities. The chamber is sealed again and then 1/5th of the products flushed out by an appropriate concentration of alcohol. The alcohol displaces the DNA strands from the magnetic beads. The sequencing reaction products can be taken out of the chip and electrokinetically injected into the

AB3730 detectors.

Microfluidic Metering and Pumping

This first design as proposed was quite ambitious. It was desirable to the break up the individual DNA reactions involved and study them individually. But before that, the first practical stage of the project was to test the microfluidic devices for basic operations such as valve opening/closing, microfluidic metering and pumping.

Figure

2.2

Microfluidic Device based on Design 1

The device shown above in Figure 2.2 was designed to meter the requisite amounts of

templi-phi and the DNA template for amplification. This is the same design as proposed

in Figure 2.1. Figure 2.2 is indeed Figure 2.1 rotated counterclockwise by 90 degrees!

The channel width here is 100 ptm and the height is 10 pim. The reactants metered in the

metering region (as in Figure 2.1) would be forced into the circular peristaltic mixer for

mixing followed by amplification. The metering region is zoomed-in and shown in

Figure 2.3.

The valves 1,2,3,4 were used to meter in the template. The volume 1-4 as shown in the

figure accommodates approximately 0.4 nI of Template. The valves 4,5,6,7 were used to

meter templi-phi. The volume 4-7 as shown accommodates about 2.5 nl of TempliPhi.

Figure 2.3 Metering Region in Design 1

The aforementioned volumes were not arrived at arbitrarily. As analyzed by Joe Graham

(Broad Institute) in Table 1.1, on a microfluidic platform it is indeed unnecessary to first

do a highly concentrated DNA T-phi reaction and then take a miniscule portion of it

discarding the rest. In the 384 well plate reaction, this strategy becomes necessary

because the amplification reaction does not proceed well until there is a minimal

concentration of reactants. It was shown in Table 1.1 that discarding the 2 stage dilutions,

we can afford to manage with only 8 nl of template and about

50

nl of TempliPhi.

Figure 2.4. Cross-Injection across valves 5,6 bounded by valves 4,7

Further, it was observed that such a reaction still produces excess sequencing product.

An analysis by J. Meldrim[2] showed that only 2% of the sequencing product may be

enough to produce decent reads on the 3730 detector. We decided run the reaction with

5%

of the volumes suggested in the scale-down of Table 1. Thus the volumes of

Template and Templi-phi required scale down to 0.4 nl and 2.4 nl respectively. This

linear scale-down is a heavy approximation and suitable experimentation is needed to

find out if this is applicable.

Here the methodology of cross-injection is defined. Lets take the case of valves 4-7 for

metering Templi-phi shown in green in Figure 2.4. Template, as shown in orange, has

already been metered. Valves 4,7 should be closed prior to the operation. Now, valves

5,6

- iiiimi ''I I I I m.iiii... 11111111 111 I Iii -. - 11T - - -

-Peristaltic Pump Variable Metering

This situation is exactly shown in Figure 2.4. As soon as the fluid comes out of the outlet

(this operation can be suitably 'timed' via Labview controls), valve 6 is closed. Valve

5

is

still open so that fluid completely fills the recesses of the channel and forces out the air

through the ends of the valved-off regions. After it is observed through the microscope

that there are no more vacant spaces in the channel, valve

5

can be closed off as well.

Figure 2.6 Microfluidic Device based on Design 2 - Metering valves are numbered

Now valve 4 separating Template and Templi-phi is opened and the mixture is forced

into the circular chamber through dilution water. This amplification mix occupies

/

of

the circular chamber.

It was noted, however, that such a design as in Figures 2.1-2.4 does not allow one to vary

the volume of Templi-phi between successive experiments. To see the variation of

amplified product with the amount of Templi-phi used, a new chip would have to be

designed with a different channel length between valves 4,7. It was quite easy, however,

to vary the amount of input DNA template in the reaction since input DNA is available in

a variety of concentrations. But Templi-phi comes in a standard concentration and so we

decided to design a chip wherein variable volumes of Templi-phi could be metered.

The corresponding graphic design 2 is shown in Figure 2.5 and Figure 2.6. Here it is

possible to meter variable volumes of Templi-phi in increments of 0.5 nl. The smallest

volume available for Templi-phi metering is 0.5 nl between valves 3,4 with inlet and

outlet bounded by valves 1,2 respectively. The next larger volume is 1 nl between valves

3,5 followed by a volume of 1.5 nl between valves 3,6 and so on.

Once Templi-phi has been metered as needed, it can be forced into the rotary mixer

chamber by pushing the diluted Template into the channel. This operation is illustrated in

Figure

2.7

and Figure 2.8. In figure 2.7, 1.5 nl of Templi-phi, in green, has been metered

between valves 3 and 6. In Figure 2.8, Templi-phi is forced ahead into the rotary mixer

by the appropriately diluted template, in yellow, into the rotary mixer. The templi-phi,

template mix has been forced to fill only a portion of the rotary mixer in Figure 2.8. This

is done just as a demonstration of the valve and metering controls. In a practical operating

situation the mixture will fill the complete rotary mixture. The template will have to be

appropriately diluted so that the required amount of template DNA is available for

amplification reaction.

Figure 2.8 Metered templi-phi forced into the rotary mixer by the template

.. -- -_- ----- o;- - - _-I--- _

opposed to 50 nl earlier on. The radius of the annular chamber is 0.8 mm as compared to 4 mm in design 1. The width and height of the channels is 100 ptm and 20 pim respectively. The lesser diameter of the annular chamber allows much faster peristaltic pumping.

As the valving and cross-injection aspects of microfluidics had already been investigated in design 1, the primary application of design 2 was to test peristaltic pumping as demonstrated by Unger et. al.[1]. Complete mixing of the reactants in this design is completed in less than a minute. Figure 2.9 shows the scenario after the mixing of three colored dyes - orange, yellow and green.

References

[1] Unger MA, Chou HP, Thorsen T, Scherer A, Quake SR, "Monolithic Microfabricated

Valves and Pumps by Multilayer Soft Lithography", Science 288: 113-116 (2000)

[2] Meldrim, J., Limit of Detection, Technology & Development at the Broad Institute Presentation, 2004

Chapter 3

Amplification

In order to systematically work towards the integrated project goals the final task was divided into individually manageable sub-tasks. As Templi-phi amplification was the first step in the sample preparation protocols, initial efforts focused on the development of a microfluidic chip to automate the amplification process. Chip design 2 was used as the

foundation for this work.

Amplification Mechanism of TempliPhi

The amplification technique used at the Broad Institute is called the Rolling Circle Amplification (RCA). This differs from the conventional Polymerase Chain Reaction (PCR) amplification method, which cycles through a temperature range for each base extension, in that it is carried out isothermally (300 C). This greatly simplifies the

RCA eliminates the requirement for extended bacterial growth prior to sequencing and saves laboratory personnel hands-on time by eliminating the centrifugation and transfer steps required by older preparatory methods. Additionally, costly purification filters and columns are not necessary, as amplified product can be added directly to a sequencing reaction. Starting material can be any circular template from a colony, culture, glycerol stock or plaque.

TempliPhi 10,000

TempliPhi 100/500

Time (hours) Time (hours)

Figure 3.1 Templi-phi kinetics for the Standard Manufacturer kits

Before RCA, the conventional method for sequencing template production was bacterial culture - a labor intensive multi-step procedure that takes up to 24 h and produces templates of varying quality and quantity [1]. This deficiency is overcome in RCA. The biggest advantage of RCA is really that the amplification product can be directly

sequenced, eliminating the need for costly commercial purification columns and plates.

Templi-phi is available generally in two varieties, TempliPhi 500 and TempliPhi 10000

nanoliter scale. TempliPhi 10000 kit comes with all the components mixed together in proportions presumably optimized on the microliter scale.

As Figure 3.1 from the manufacturer's catalogues demonstrates, both of the product kits (Templi 500 and Templi-phi 10000) can achieve sufficient amplification within 4-6 hours. However, at the Broad Institute, the amplification routinely is done using only the TempliPhi 10000 kit and the process is designed to be completed within 16 hours. The idea in this section will be to find out how this kinetics scales down when the reaction is performed at nanoliter scale in the microfluidic environment.

Rolling circle amplification (RCA) is a technique used to amplify circularized DNA with a sequence specific primer and a strand displacing DNA polymerase, producing a repeating linear construct. The polymerase, TempliPhi from the bacteriophage Phi29, processively creates a copy of the circular plasmid DNA template utilizing random hexamer primers. RCA was discovered owing to the observation that replicating phage

DNA in infected bacteria was larger than in phage particles because of the formation of

concatamers of genomic phage DNA. Since phage DNA is circular, tandem repeats are produced by rolling circle replication (RCR), in which the phage DNA polymerase replicates and subsequently displaces the newly made strand.

The reaction begins with denatured single stranded circular DNA

Phi29 DNA polymerase binds the primers

Polymerase replication

C)

Most DNA polymerases will

stop here, unable to displace

the newly replicated downstream

I

Phi29 DNA polymerase is unique,

it begins strand displacement with no loss of replication speed at all

Strand displacement

The newly displa There is no need single stranded D

new DNA is a tem

ced strands are single stranded. More random primers for thermal cycling to create bind the new displaced strands

NA (as with PCR), and this and ar xtended by polymera

Eventually all of the nucleotides are depleted and the reaction stops. By this time the enzyme will have made 4-5 pg of double stranded, tandemly repeated copies of the circle.

This DNA does not need to purified,

and can be added directly to a sequencing reaction

Figure 3.2 Mechanism of Rolling Circle Amplification

Random hexamers bind to the denatured circular template allowing Phi29 DNA polymerase to initiate multiple amplification events (Figure 3.2). The inherent strand-displacement activity of the enzyme displaces the 5'-ends of downstream strands. As

DNA synthesis and displacement continue, the enzyme produces single-stranded,

complementary concatamers of the circular template (Figure 3.2). Priming and polymerization directed by the displaced strands produces double-stranded DNA.

Nucleotides in the TempliPhi premix fuel the reaction to produce as much as 5 pig of

DNA in the pl scale reaction in a 384 well plate. Amersham Biosciences catalogues claim

this can be exceeded and DNA amplification completed from much lower starting template amounts.

The rate of the strand synthesis by Phi29 DNA polymerase is claimed to be approximately 50 nucleotides/second, due in part to the enzyme's high processivity. Phi29 DNA polymerase is able to incorporate greater than 70,000 nucleotides during a single binding event without the aid of accessory proteins.

The microfluidic experiments outlined in this thesis are designed to test the lower limits of input template that can be used as claimed by Amersham Biosciences. A parametric study was carried out in the microdevices, varying input template and Templi-Phi polymerase concentrations as well as the total reaction time. In the process of protocol development, the microlfuidic device underwent several design modifications to control issues like sample evaporations and biofouling of the microchannel walls.

Fluorescence Measurement Methodology

The first issue towards the above mentioned goals was to find out an effective way of quantifying the on-chip Templi-phi reaction. An early option investigated was to take the amplified DNA out of the chip and determine the extent of amplification by gel-electrophoresis. However, the quantity produced on chip was below the threshold of standard quantification by gel-electrophoresis.

Preliminary calculations from Table 1.1 indicated that 0.6 ng of amplified DNA product may be required to feed into the subsequent DNA sequencing reaction. Off-chip quantification of the amplification product, even if feasible, introduces the possilibty of sample loss or contamination that compromises the sequencing reaction. To realize the true utility of a microfluidic platform, it is desirable to perform as many operations as

possible on the chip itself. Hence, the idea of using DNA quantification fluorescent dyes was employed. There were several candidates like Pico Green, SYBR Green, Oligreen, Ethidium Bromide, Hoechst 33258 etc. Since 80% of the product generated by RCA is double-stranded, single-stranded detection dyes like Oligreen cannot be used. The dye should really be dsDNA specific and ideally show no fluorescent enhancement in the presence of ssDNA. Further, we need sensitivity of detection to small quantities of DNA at the scale of -1 ng since we need to calibrate the extent of DNA amplification with respect to our input parameters (input DNA template Templi-Phi concentrations, amplification time).

It was found that PicoGreen was best suited to the application at hand. PicoGreen is an intercalating agent, and is inserted between the individual DNA strands. The concentration of intercalated PicoGreen is linear with respect to the number of base pairs of DNA, and the fluorescence emission of the dye can be used to obtain quantitative measurements of DNA concentration. The fluorescent enhancement of this dye is >1000 fold upon binding to dsDNA with excitation around 495 nm and emission around 520 nm.

Unlike Hoechst 33258 and some other dues, PicoGreen fluorescence intensity is the same upon binding to poly(dA).poly(dT) and poly(dG).poly(dC) homopolymers. This is important for providing a dependable measure of DNA concentration. The linear concentration range for DNA quantitation extends over four orders of magnitude - 25 pg/ml to 1 ptg/ml - with a single dye concentration [2].

However, in our case, it is expected that the post-amplification concentrations in the microfluidic chamber might be of the order of 100 ptg/ml. This would not only lie outside the linear range of PicoGreen but also out of the routinely tested range of most of the dyes. It was interesting to find out whether PicoGreen would provide a sufficient

curve was indeed found out and it provided good variation over the pertinent range of quantitation.

Firstly, a set of titration experiments was performed to calibrate fluorescence emission in the microfluidic channels. Certain issues had to be handled well here. There is always some variability in the microfabrication procedure varying the height of microfluidic channels in spite of using the same exposure mask in photolithography. To ensure that the set of channels used for calibration and the set of channels used for successive real experiments had the same height dimensions, a profilometer was used to measure channel-channel variations between different devices. Final devices selected for experimentation has height variations of less than one micron vs. an average channel height of 20 microns.

The fluorescent images were captured from a fluorescence microscope (Nikon TE -2000, 20X Objective). Figure 3.3 shows a snapshot of 10 nM concentration of DNA in the channel. The fluorescent values were obtained using ImageQuant software. It should be noted here that the fluorescence observed depends not only on the channel depth but also the exposure settings of the microscope. For Fig.3.3, the settings were Gain = 16 and Capture Time = 31 ims. It was necessary to adopt the same exposure settings in real experimentation as in calibration.

In order to increase the amount of final product obtained, the height of the channels in design 2 was increased to 20 [tm (from 10 ptm in Design 1). This corresponds to a volume of 10 nl in the peristaltic mixer as mentioned earlier. A final amplified product of 1 ng corresponds to 55 nM of DNA in a 10 n1 reaction. Hence we would expect much brighter fluorescence compared to Figure 3.3, which is only 5 nM, if the Templi-phi reaction proceeds successfully.

Figure 3.3 PicoGreen Fluorescence image for IOnM of pUC19 DNA in the microchannel.

It was subsequently decided that PicoGreen would not be used as a part of the Templi-phi

reaction mix as the dye intercalates between the grooves of the DNA and hence might

hamper the activity of the Phi 29 DNA polymerase. Hence, PicoGreen was added only

after the amplification reaction was over.

Referring to Figure 3.4, once the reaction was completed, valves 2,3 were opened while

keeping valves 1,4 closed. PicoGreen was forced through valves 1,4 flushing some of the

amplification product outside the reaction chamber. The region 1-4 however comprises

only

/7th of the reaction volume and the remaining

/7th of the reaction product is still

available to mix with PicoGreen reagent. Valves 1,4 are now closed and valves 2,3

It should be noted that valves 1,4 serve the dual purpose of acting as gates to allow only

a specific amount of amplified product to be replaced with PicoGreen and also as two of

the three valves required for peristaltic pumping.

Commercially available 1-X PicoGreen reagent is very concentrated and is generally

diluted before using for DNA quantitation. Here 1/10-X PcioGreen is used. This strength

is arrived at by linearly scaling the amount of dye required to detect DNA on the order of

5

ng.

t

PicoGreen In

PicoGreen Out

-r

Figure 3.4 Valves 1,4 act as gates to allow only a specific amount of amplified product to be replaced with PicoGreen reagent. They are also 2 of the 3 valves required for peristaltic pumping!

I

I

2

I1

13

Amplification Trials

Appendix III shows all amplification reaction trials in tabular form.

The first trials of Templi-phi reaction on chip were performed using design 2 (Figure 2.5) which had a reaction volume of 10 nl. Relying on preliminary scale-down calculations from Table 1.1 and some guess work, the first reaction mix tried was 2 nl 10000 Templi-phi + 8 nl 1 nM template with a reaction time of 30 min.

The amount of Templi-phi was scaled down by three orders of magnitude from the 3 pl used in the 384 well plate reactions. According to a linear scaling analysis, the final amplified product was predicted to be 1-2 ng down by three orders of magnitude from the

2-2.5 ptg produced in the 384 well plate reactions.

The time required for reaction was also scaled down from 4-16 hours to 30 min. It was argued that the time needed for the reaction to finish was really the time for the enzyme to incorporate all the nucleotides. Since the amount of nucleotides and template has been scaled down by three orders of magnitude, the time should also scale down drastically. Accordingly the time was scaled down by one order of magnitude, this being an initial guess only. A lesser reaction time could have been assumed but a conservative estimate was preferred in the initial set of experiments. The template used was pUC 19, which is generally provided as a control in Templi-phi reaction kits.

Thirty minutes after the reactants were mixed on-chip, the product was tested within PicoGreen in the way outlined in the preceding section. However, no change in fluorescence was observed, suggesting that the reaction was not taking place.

any effects of background fluorescence and also of the initially present double stranded template were accounted for.

Subsequently, the possible failure scenarios were examined. The amount of starting template used was 36 pg and it could be argued that it was too low for any significant amplification to take place. However, in the catalogues of Amersham Biosciences for TempliPhi, it was reported that robust amplification was observed with as low as 1 pg of starting plasmid DNA [3].

Another pertinent idea was that the binding of hexamers, present in TempliPhi mix, with the template is thermodynamically more favorable if the template is single stranded rather than double stranded. The template is indeed denatured in the 384 well plate reactions to allow the vector DNA to escape form the E. Coli host cells without lysing the cells, producing single stranded DNA in the process. Hence, it was decided to denature the template off-chip before proceeding with the on-chip reaction.

Further, a careful revision of enzyme kinetics tells us that the time required for reaction to complete may not scale down as drastically as previously expected.

dS

dt

S

where S is the concentration of the substrate at any time. The above relation indicates that the time taken depends on the fractional change in concentration of the substrate concentration. Accordingly, even when the amount of substrate and enzyme is scaled down tremendously, the reaction time may not decrease at all since the fractional change in concentration of the substrate required to complete a portion of the reaction still remains the same in the scaled-down reaction as it was in the original reaction. Keeping

this analysis in mind, it was also decided to run the experiments for longer periods of time identical to the 384 well plate reactions.

Hence, several reactions were tried where the template was denatured prior to the reaction and with times varying from 2 hours to 16 hours. However, as the reaction time went into hours from minutes, it was observed that the reaction mix evaporated almost completely even at room temperature. In spite of the valves remaining closed throughout the reaction, the reaction mix evaporated invariably within an hour from the onset of the reaction.

It was recognized then that PDMS is gas-permeable and so it easily allows water vapor to escape through the pores in the polymer matrix. Several procedures have been used by researchers to address this issue. The channels can be coated with suitable chemicals to inhibit evaporation. However, here one has to careful in choosing the chemical regarding its affinity with DNA and enzymes. A three-layer PDMS device can be constructed instead of a two-layer device wherein the topmost layer has a network of water jackets. Using water jackets above the reaction channels provides a saturated environment in the polymer pores and so evaporation is curbed.

However, in the present device it is much easier to incorporate these water jackets in the control layer, which lies about 20 pm above the flow layer. Figure 3.5 shows the water jackets and other control layer channels in red around the black reaction chamber. With

Water Jackets to counter Evaporation

Figure 3.5. Spiral Water Jackets around the Circular Reaction Chamber to provide a Saturated Environment

With the modified design as shown in Figure

3.5,

longer reactions were performed.

Appendix III shows all the amplification reaction trials in tabular form. Still only 2-3 fold

amplification was observed.

Further troubleshooting led to the hypothesis that the enzyme availability was limited in

the microfluidic devices due to partitioning of enzyme into the PDMS or adhesion to the

PDMS microchannel walls. To check how much the PDMS surface can impede the action

of the enzyme, several reactions were performed in the 384 well plate format with some

small pieces of PDMS submerged in the reaction mix. However, there was no

considerable decrease in the amount of amplified product and subsequent generated

sequence.

In spite of negligible effect of incorporating PDMS pieces into the 384 well plate

reaction, nothing can be inferred about the corresponding behavior at the micro-scale as

the surface area to volume ratio increases tremendously in microfluidic devices. Hence,

One possibility could be that the amount of enzyme scaled down by three orders of magnitude for the micro-scale reaction was just not significant enough to catalyze the reaction. Another more feasible scenario was that the PDMS channel surfaces may lead to biofouling and hence the leave very little enzyme available in solution to participate in the reaction.

Hence, a decision was made to use TempliPhi 500 kit instead of the TempliPhi 10000 kit. This was because the TempliPhi 500 kit provides the enzyme, dNPTs and the hexamers/reaction buffer separately as opposed to the TempliPhi 10000 kit, which bunches the three together in unknown concentrations. Hence, we would have the flexibility to play with the amounts the enzyme and other constituents.

In the next set of reactions, the Phi-29 enzyme was used in excess as compared to the manufacturer's specifications. Also, the template was denatured off-chip and pre-annealed with the reaction buffer mixture containing the random hexamers. However, no amplification was observed.

Number of Spiral Water Jackets is

Number of Spiral Water Jackets is

increased to enable longer Reactions.

Metering is no longer performed on-chip.

Figure 3.6 Microfluidic Layout around the Circular Mixer for Design 3.

As the amplification protocols failed on the chip in spite of the troubleshooting efforts, one final hypothesis was explored. Given the complexity of the amplification process, utilizing multiple substrates with reaction kinetics that are not first order, perhaps the scalability of the reaction collapses as critically small reaction volumes owing to a combination of the aforementioned problems (i.e. substrate or enzyme concentrations) or subtle changes in the buffer composition due to evaporation or osmotic equilibration. To test this hypothesis, we carried out a final set of experiments with scaled-up reaction volumes on-chip.

The total reaction volume in Design 3 is 40 nl. The width of flow channels is 200 pm and the height is 20 pm. Further the radius of the annular chamber is increased to 3.2 mm from 1.6 mm in Design 2. Complete mixing is still observed to happen within 90 seconds in this design. Further, the number of spiral jackets is greatly increased as compared to

Figure 3.5. These water jackets make possible reaction times up to 24 hours and longer

without any significant evaporation.

Figure 3.7 Successful amplification gives a strong fluorescence signal - Row 9, Appendix III

The reaction mix for this 40 nl reaction can be seen in row 9 in Appendix III. It

comprises of 24 nl TempliPhi 10000 and 15.5 nl TempliPhi 500 (6 nl dNTPs, 5.5 nl

reaction buffer and 4 nl enzyme). For the first time, a strong fluorescence signal was

obtained from the PicoGreen indicating a successful reaction. From a starting template

amount of 45 pg and a reaction time of 16 hours, a strong amplification of 80 fold was

observed leading to an amplified product of 3.5 ng. The corresponding fluorescence

image is shown in Figure 3.7.

The amplified product from the above reaction was purged off the chip, diluted to 1.3 g1

and mixed with 0.75 p1 of Big Dye mix to yield 2.05 pl of sequencing reaction mix. This