Microfluidic Interfaces for Mass Spectrometry: Methods and Applications

by

HAO YANG

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy – Analytical Chemistry

Department of Chemistry University of Toronto

© Copyright by Hao Yang (2011)

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Microfluidic Interfaces for Mass Spectrometry: Methods and

Applications

Hao Yang

Doctor of Philosophy – Analytical Chemistry

Department of Chemistry

University of Toronto

2011

Abstract

Since the introduction of electrospray ionization (ESI) and matrix assisted laser

desorption ionization (MALDI), there has been an unprecedented growth of biomolecule analysis

using mass spectrometry (MS). One of the most popular applications for mass spectrometry is

the field of proteomics, which has emerged as the next scientific challenge in the post-genome

era. One critical step in proteomic analysis is sample preparation, a major bottleneck that is

attributed to many time consuming and labor-intensive steps involved. Microfluidics can play an

important role in proteome sample preparation due to its ability to handle small volumes of

sample and reagent, and its capability to integrate multiple processes on a single chip with the

potential for high-throughput analysis. However, to utilize microfluidic systems for proteome

analysis, an efficient interface between microfluidic chip and mass spectrometry is required. This

thesis presents several methods for coupling of microfluidic chips with ESI-MS and MALDI-

MS.

III

Three microfluidic-ESI interfaces were developed. The first interface involves fabricating

a polymer based microchannel at the rectangular corners of the glass substrates using a single

photolithography step. The second interface was build upon the previous interface in which a

digital microfluidic platform was integrated with the microchannel in a “top-down” format. The

integrated microfluidic system was used for inline quantification of amino acids in dried blood

spots that have been processed by digital microfluidics. The third interface was formed by

sandwiching a pulled glass capillary emitter between two digital microfluidic substrates. This

method is a simpler and more direct coupling of digital microfluidics with ESI-MS as compared

to the method used for second interface. Finally, a strategy using a removable plastic “skin” was

developed to interface digital microfluidics with MALDI-MS for offline sample analysis. We

demonstrated the utility of this format by implementing on-chip protein digestion on

immobilized enzyme depots.

IV

Acknowledgments

I would like to give my deepest gratitude to my supervisor, Dr. Aaron Wheeler. He gave

me the opportunity to join the group at University of Toronto and work on the exciting

microfluidic interface project. Without his constant guidance and valuable advices, I would not

have accomplished this goal.

I would also like to thank my PhD committee members: Professor Rebecca Jockusch,

Professor Mark Nitz, and Professor Ulrich Krull. Your constructive feedbacks during annual

committee meetings have steered these projects and my development as a researcher in the

positive direction. Likewise, I would like to acknowledge Professor Kagen Kerman for

participating in my comprehensive oral examination and my thesis defense.

I am grateful to many previous members of my group: Dr. Irena Barbulovic-Nad, Dr.

Beth Miller, Dr. Jianhua Qin, Dr. Edmond Young, and Uvaraj Uddayasankar. I was very

fortunate to work with them in several occasions and they have taught me numerous lessons

from their own experience. I want to thank Dr. Sergio Freire, in particular, for showing me

around as a mentor, and getting acquainted with the fabrication processes for my nanoESI chip

interface. Also Dr. Mohamed Abdelgawad and Dr. Noha Mousa, I enjoyed working with them

on several different projects. They were not only good companions, but also wonderful friends

outside the lab.

I am also thankful to all the current members of my group: Dr. Dario Bogojevic, Dr.

Lindsey Fiddes, Dr. Yan Gao, Dr. Michael Watson, Sam Au, Irwin Eydelnant, Ryan Fobel, Mais

Jebrail, Andrea Kirby, Nelson Lafreniere, Vivienne Luk, Jared Mudrik, Alphonsus Ng, Steve

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Shih and Suthan Srigunapalan. You have been a special group to work with and I wish you all

success in what will certainly be bright futures.

I would like to acknowledge Dr. Henry Lee and Yimin Zhou for their dedication to the

ECTI cleanroom. Their efforts in maintaining a working facility has been crucial for this success

work. Likewise, I would like to thank the entire crews of the machine shop in the department.

Without their magical hands, I would not be able to accomplish my goals that require

modification of instruments and building custom setups.

Last but not least, I would like to thank my parents, Frank and DaFang for their supports

and encouragements throughout my career choices. I have been blessed with their endless love.

Over the years and through lots of sacrifices, they have laid the ground work for me to reach

where I am now. I hope I have realized their dreams and made them proud.

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Table of Contents

Acknowledgments......................................................................................................................... IV Table of Contents .......................................................................................................................... VI Overview of Chapters ................................................................................................................... IX Overview of Author Contributions .............................................................................................. XII List of Tables ............................................................................................................................... XV List of Figures ............................................................................................................................ XVI List of Abbreviations ............................................................................................................... XVIII Chapter 1 Review of Microfluidic Interfaces for Mass Spectrometry ...........................................1

1.1 Mass Spectrometry ...............................................................................................................1 1.2.1 Ionization Methods ..................................................................................................1 1.2.2 Mass Spectrometry for Proteomics ..........................................................................3

1.2 Microfluidics ........................................................................................................................6 1.2.1 Channel Microfluidics .............................................................................................6 1.2.2 Digital Microfluidics ..............................................................................................12

1.3 Microfluidic Interfaces for Mass Spectrometry .................................................................22 1.3.1 Microfluidic-ESI-MS Interfaces ............................................................................22 1.3.2 Microfluidic-MALDI-MS Interfaces .....................................................................28

1.4 Conclusion .........................................................................................................................32 Chapter 2 A Practical Interface for Microfluidics and Nanoelectrospray Mass Spectrometry ....33

2.1 Introduction ........................................................................................................................34 2.2 Materials and Methods .......................................................................................................36

2.2.1 Reagents and Materials ..........................................................................................36 2.2.2 Device Fabrication .................................................................................................37 2.2.3 Device Operation ...................................................................................................38 2.2.4 Mass Spectrometry .................................................................................................39

2.3 Results and Discussion ......................................................................................................41 2.3.1 Device Fabrication and Operation .........................................................................41 2.3.2 Device Performance ...............................................................................................42 2.3.3 Mass Spectrometry Performance ...........................................................................45

2.4 Conclusion .........................................................................................................................49 Chapter 3 A Digital Microfluidic Method for Amino Acid Quantification in Blood ..................50

3.1 Introduction ........................................................................................................................51 3.2 Materials and Methods .......................................................................................................54

3.2.1 Study Subjects ........................................................................................................54 3.2.2 Reagents and Materials ..........................................................................................54 3.2.3 Reagents and Materials for Device Fabrication .....................................................55 3.2.4 DMF Device Fabrication and Operation ................................................................55 3.2.5 Hybrid Device Fabrication and Operation .............................................................58

VII

3.2.6 Non-DMF Sample Processing and Analysis..........................................................59 3.2.7 DMF-Driven Sample Processing ...........................................................................59 3.2.8 DMF-Driven Sample Analysis ..............................................................................60 3.2.9 DMF-Driven Sample Recovery Evaluation ...........................................................61

3.3 Results ................................................................................................................................63 3.3.1 DMF-Driven Method 1 ..........................................................................................64 3.3.2 DMF-Driven Method 2 ..........................................................................................68 3.3.3 DMF-Driven Method 3 ..........................................................................................70

3.4 Discussion ..........................................................................................................................73 3.5 Conclusions ........................................................................................................................76

Chapter 4 Digital Microfluidic Electrospray Ionization Interface for Succinylacetone

Analysis in Dried Blood ............................................................................................................77 4.1 Introduction ........................................................................................................................78 4.2 Materials and Methods .......................................................................................................80

4.2.1 Study Subjects ........................................................................................................80 4.2.2 Reagents and Materials ..........................................................................................80 4.2.3 DMF Device Fabrication .......................................................................................80 4.2.4 DMF Device Operation ..........................................................................................82 4.2.5 DMF-ESI-MS Interface .........................................................................................84 4.2.6 DMF-driven SA Processing ...................................................................................84 4.2.7 SA Quantification by NanoESI-MS/MS ................................................................85

4.3 Results and Discussions .....................................................................................................86 4.3.1 DMF Fabrication and Operation ............................................................................86 4.3.2 DMF-ESI-MS Interface .........................................................................................87 4.3.3 DMF-driven SA Processing and Quantification ....................................................89

4.4 Conclusion .........................................................................................................................93 Chapter 5 A World-to-Chip Interface for Digital Microfluidics ..................................................94

5.1 Introduction ........................................................................................................................95 5.2 Materials and Methods .......................................................................................................97

5.2.1 Reagents and Chemicals ........................................................................................97 5.2.2 Device Fabrication and Operation .........................................................................98 5.2.3 Mass Spectrometry ..............................................................................................100 5.2.4 Fluorescence ........................................................................................................101

5.3 Results and Discussion ....................................................................................................103 5.3.1 Protein Adsorption and Cross Contamination .....................................................103 5.3.2 Digital Microfluidic Skins ...................................................................................104 5.3.3 Skin Depot ...........................................................................................................108 5.3.4 Skin Depot Stability .............................................................................................112

5.4 Conclusions ......................................................................................................................114 Concluding Remarks and Future Potential .............................................................................115

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Appendix 1 Digital Microfluidics for Solid Phase Extraction ....................................................118 A1.1 Introduction ..................................................................................................................119 A1.2 Materials and Methods .................................................................................................121

A1.2.1 Reagents and Materials .......................................................................................121 A1.2.2 Device Fabrication and Operation ......................................................................122 A1.2.3 PPM Formation and Characterization .................................................................125 A1.2.4 Standard DMF-SPE Process ...............................................................................126 A1.2.5 DMF-SPE Process Optimization ........................................................................126 A1.2.6 DMF-SPE Extraction Efficiency ........................................................................127 A1.2.7 DMF-SPE for Preparative Sample Cleanup .......................................................128 A1.2.8 DMF-SPE for Sample Concentration .................................................................129

A1.3 Results and Discussion .................................................................................................130 A1.3.1 PPM Formation and Characterization .................................................................130 A1.3.2 DMF-SPE ............................................................................................................132 A1.3.3 DMF-SPE for Preparative Analysis ....................................................................134 A1.3.4 DMF-SPE for Sample Concentration .................................................................136

A1.4 Conclusion ....................................................................................................................138 References ...................................................................................................................................139

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Overview of Chapters

This thesis describes various microfluidic interfaces for mass spectrometric applications

that we developed while working in the Wheeler Group. These interfaces have been categorized

into two sections: Microfluidic-electrospray ionization mass spectrometry (ESI-MS) interfaces

(Chapter 2, 3, and 4), and microfluidic-matrix assisted laser desorption ionization mass

spectrometry (MALDI-MS) interfaces (Chapter 5). Here, for the benefit of the reader, I have

provided a brief outline of each chapter’s contents.

Chapter one provides a literature review of various existing microfluidic interfaces for

mass spectrometry, in three sections. First, I present a brief overview of two popular ionization

techniques in mass spectrometry (MS), with a short review of their uses in MS techniques for

proteomics. Second, I introduce microfluidic technologies as a useful tool for proteomic sample

preparation and analysis. Third, I review several strategies that have been developed to couple

these ionization techniques with microfluidic devices.

Chapter two describes a new method for fabricating nanospray ionization tips for mass

spectrometry. The tip was fabricated from ultra-thin glass substrates and an inert polymer,

Parylene C, using a single photolithography step. The emitter is formed contiguously with the

microchannel such that there is no dead volume. Various analytes ranging from synthetic

polymers, peptides, and even nucleic acids were sprayed in these devices, and the performance

was comparable to conventional pulled-glass capillary emitters as well as the Agilent HPLC

ChipTM.

Chapter three describes an integrated microfluidic system for processing newborn blood

samples for analysis by tandem MS (MS/MS). The method is capable of processing two different

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types of blood samples: blood directly spotted onto the chip, and dried blood spots on filter

paper. In addition, some devices were integrated with a microchannel emitter for inline analysis

of the processed blood samples by tandem MS. The method is fast, robust, precise, and is

capable of quantifying analytes in blood samples associated with common congenital disorders

such as homocystinuria, phenylketonuria, and tyrosinemia.

Chapter four represents work in progress, and it describes a very straightforward digital

microfluidic interface to MS for analysis. In this method, a pulled glass nanoelectrospray emitter

is aligned between the two plates of a digital microfluidic device with an inter-plate spacing that

is equal to the outer diameter of the glass capillary. Sample droplets were processed by DMF and

then actuated to the capillary interface such that the glass emitter is filled by capillary action. A

nanoelectrospray was generated by application of high voltage to the counter electrode of the

DMF setup. The system was validated by application to automated extraction of succinylacetone

(SA) from dried blood spot (DBS) with inline quantification of SA by MS/MS.

Chapter five introduces a new strategy for digital microfluidics, in which a removable

plastic “skin” is used to (a) eliminate cross-contamination, (b) bridge the world-to-chip interface,

and (c) interface with MALDI-MS. A new skin can be positioned on the device at the beginning

of each experiment, and may be modified to contain surface-adhered reagents. The utility of this

format was demonstrated by implementing on-chip protein digestion on immobilized enzyme

depots.

The Appendix describes an extra project that did not fit the overall narrative of this thesis.

It introduces the marriage of two technologies: DMF, a technique in which droplets are

manipulated by application of electrostatic forces on an array of electrodes coated by an

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insulator, and porous polymer monoliths (PPMs), a class of materials that is popular for use for

solid phase extraction and chromatography. In this work, PPM discs were formed in situ by

dispensing and manipulating droplets of monomer solutions to designated spots on a DMF

device, followed by UV-initiated polymerization. PPM discs formed in this manner were used to

develop a digital microfluidic solid phase extraction (DMF-SPE) method, in which PPM discs

were activated and equilibrated, samples were loaded, PPM discs were washed, and the samples

were eluted, all using microliter droplets of samples and reagents.

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Overview of Author Contributions

During the course of my research, I was fortunate to work with a number of collaborators

in and out of the Wheeler group, many of whom are co-authors on the journal papers that have

been published or are in review. Here I outline the contributions each person made towards the

work presented herein.

In the work described in Chapter two, Dr. Sergio Freire, (then post doc) developed the

fabrication protocol and drew the photomasks for printing. I had the ideas of using thin

microscope glass cover slips to reduce the wetting surface, and for fabricating the microchannel

at the corner of the device to mimic the tapering geometry of conventional microfabricated

emitters. Sergio and I worked together to fabricate the devices in the cleanroom and assemble the

devices with commercially available fittings. Likewise, we worked together to perform the

experiments and prepare the figures for the manuscript. This work was published in

Electrophoresis (Freire, S.L.S.; Yang, H.; Wheeler, A.R. Electrophoresis, 2008, 29 (9), 1836-

1843) on which I am a co-author.

The work described in Chapter three was carried out in collaboration with the Newborn

Screening Ontario (NSO) facility at the Children's Hospital of Eastern Ontario in Ottawa,

Ontario. Several members from NSO participated in this project. Specifically, Christine

McRoberts (technician), Lawrence Fisher (technician), Dr. Osama Al-Dirbashi (scientist), and

Dr. Pranesh Chakraborty (director) contributed valuable suggestions and provided newborn

patient samples. Christine, Lawrence and Osama had carried out analyses using conventional

methods for comparison to the microfluidic methods developed in the Wheeler group. From the

Wheeler group, (then graduate student) Dr. Mais Jebrail designed and operated the basic device

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in which samples were processed by DMF with off-line analysis by mass spectrometry. Graduate

students Jared Mudrik and Nelson Lafrenière assisted in fabricating these devices. I designed,

fabricated, and operated the DMF devices that were integrated with a microchannel emitter for

in-line mass spectrometry analysis. Mais and I worked together to collect and analyze all of the

mass spectrometry data. A version of this chapter has been submitted and is currently under

review. Mais and I are co-first authors of this manuscript.

In the experiments described in Chapter four, I worked with graduate student Steve Shih

who developed the feedback-controlled electronic interface and LabView software algorithms

for automated droplet movement. Mais Jebrail contributed helpful discussions and assisted with

high-definition video recording of the actuation systems. I fabricated the DMF devices and came

up the idea of sandwiching the glass emitter in between the two DMF substrates. I also designed

the extraction protocol for succinylacetone analysis in blood, and collected and analyzed all of

the mass spectrometry data. I will be the first co-author on a manuscript describing this work,

which is in preparation.

The initial idea behind the work described in Chapter five originated from (then graduate

student) Dr. Irena Barbulovic-Nad. Graduate student Vivienne Luk contributed by generating

fluorescent images of surfaces that were fouled by non-specific adsorption of fluorescently

labeled proteins. Then graduate student Dr. Mohamed Abdelgawad suggested the use of silicone

oil as an adhesive to improve annealing of polymer coverings on DMF substrates. Mohamed also

helped to create the mask for device fabrication. I made all the devices and performed the

experiments. This work was published in Analytical Chemistry (Anal. Chem. 2009, (81), 1061-

1067); I am the first author.

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In the work described in the Appendix, Mais Jebrail designed the mask, and Jared Mudrik

helped in fabricating the devices. Jared developed the methods to form and characterized PPMs

on chip. Jared and I conducted the optimization experiments to improve the extraction efficiency

of the formed PPMs. I performed the on-chip desalting and removal of surfactant experiments. In

addition, I designed and fabricated the devices used for preconcentration experiments, and

compared the extraction efficiency of my method with commercially available Ziptips. This

work has been accepted for publication in Analytical Chemistry. I am the co-first author.

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List of Tables

Chapter 3

Table 3.1 Concentrations of amino acids in blood measured by DMF-driven method 1 ..............62 Table 3.2 % Recovery of DMF-driven method 1 measured by fluorescence and MS/MS ..........68 Table 3.3 Measured Phe concentration from DBS using DMF-driven method 2 and standard technique at NSO ...........................................................................................................................70

Chapter 4

Table 4.1 Differences between the two feedback systems .............................................................83

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List of Figures

Chapter 1

Figure 1.1 Examples of integrated microchannel microfluidics ....................................................11 Figure 1.2 The DMF paradigm ......................................................................................................13 Figure 1.3 Biological applications of DMF ..................................................................................16 Figure 1.4 Schematic of various microfluidic-ESI-MS interface methods ...................................23 Figure 1.5 Examples of microchip spraying directly from edge ..................................................25 Figure 1.6 Examples of microfabricated emitters ..........................................................................27

Chapter 2

Figure 2.1 PG-NSI chip ................................................................................................................38 Figure 2.2 PG-NSI emitter fabrication process .............................................................................40 Figure 2.3 Taylor cones formed from PG-NSI emitters ...............................................................43 Figure 2.4 Typical TIC of various tested emitters ........................................................................44 Figure 2.5 MS spectra generated from PG-NSI emitters ..............................................................46 Figure 2.6 Tandem MS spectra generated from PG-NSI emitters .................................................47 Figure 2.7 Evaluation of sensitivity of PG-NSI emitters ..............................................................48

Chapter 3

Figure 3.1 Processing blood samples for quantification of AAs by MS/MS .................................52 Figure 3.2 Three DMF-driven methods designed to quantify AAs in DBS samples ....................57 Figure 3.3 Exploded view schematic of the hybrid DMF-microchannel device in method 3 .......59 Figure 3.4 Analysis of AAs in DBS by DMF-driven method 1 ....................................................64 Figure 3.5 Met, Phe, and Tyr Calibration curves generated by DMF-driven method 1 ................66 Figure 3.6 Analysis of AAs in DBS by DMF-driven method 2 ....................................................69 Figure 3.7 Analysis of AAs in DBS by DMF-driven method 3 ....................................................72

Chapter 4

Figure 4.1 DMF-nanoESI interface ...............................................................................................81 Figure 4.2 Schematic of the feedback control system ...................................................................87 Figure 4.3 Performance of the new DMF-nanoESI interface ........................................................89 Figure 4.4 Extraction of SA in DBS samples by DMF-driven method .........................................90 Figure 4.5 Tandem MS spectra of SA and 5C13 SA ......................................................................91 Figure 4.6 SA calibration curve generated by DMF-driven method .............................................92

XVII

Chapter 5

Figure 5.1 Images and spectra demonstrating protein adsorption and cross-contamination in digital microfluidics ....................................................................................................................104 Figure 5.2 Schematic depicting the removable skin strategy .....................................................105 Figure 5.3 MALDI-MS analysis of different analytes on different skins ...................................106 Figure 5.4 Schematic and spectra demonstrating the skin depot strategy ...................................109 Figure 5.5 Multiplexed skin depot ..............................................................................................110 Figure 5.6 Skin depot stability assay ..........................................................................................113

Appendix 1

Figure A1.1 DMF device design and PPM disc ..........................................................................124 Figure A1.2 DMF-SPE protocol ..................................................................................................133 Figure A1.3 DMF-SPE optimization ...........................................................................................134 Figure A1.4 Preparative DMF-SPE .............................................................................................135 Figure A1.5 DMF-SPE for concentration ...................................................................................137

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List of Abbreviations

1. 2,5-dihydroxybenzoic acid (DHB) 2. 3-hydroxypicolinic acid (HPA) 3. 3,4,5,6,7-13C-succinylacetone (513C-SA) 4. 2D gel electrophoresis (2DGE) 5. 2D liquid chromatography (2DLC) 6. Acetonitrile (ACN) 7. Acylcarnitines (AC) 8. Alpha-cyano-4-hydroxycinnamic acid (α-CHCA) 9. Amino acid (AA) 10. Angiotensin II (Ang II) 11. Brunauer, Emmett, and Teller (BET) 12. Charge-coupled device (CCD) 13. Compact disc (CD) 14. Collision induced dissociation (CID) 15. Deionized (DI) 16. Deoxyribonucleic acid (DNA) 17. Deuterated L-Methionine (Met-d3) 18. Deuterated L-Phenylalanine (Phe-d5) 19. Deuterated L-Tyrosine (Tyr-d4) 20. Digital microfluidics (DMF) 21. Digital microfluidic electrospray ionization (DMF-ESI) 22. Digital microfluidic solid phase extraction (DMF-SPE) 23. Dried blood spot (DBS) 24. Electroosmotic flow (EOF) 25. Electrospray ionization mass spectrometry (ESI-MS) 26. Emerging Communications Technology Institute (ECTI) 27. FITC-labeled bovine serum albumin (FITC-BSA) 28. Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS) 29. Hepatorenal tyrosinemia (HT) 30. Hexamethyldisilazane (HMDS) 31. High performance liquid chromatography (HPLC) 32. Hydrazine (N2H4) 33. Hydrochloric acid (HCl) 34. Indium tin oxide (ITO) 35. Inner diameter (ID) 36. Internal standard (IS) 37. Lab-on-a-chip (LOC) 38. Lateral assay tests (LAT) 39. L-Methionine (Met) 40. L-Phenylalanine (Phe) 41. L-Tyrosine (Tyr)

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42. Mass spectrometry (MS) 43. Matrix assisted laser desorption ionization mass spectrometry (MALDI-MS) 44. Methanol (MeOH) 45. Microchannel electrochromatography (MEC) 46. Microelectromechanical systems (MEMS) 47. Micro total analytical systems (µTAS) 48. Multilayer soft lithography (MSL) 49. Molecular weight search (MOWSE) 50. Nanoelectrospray ionization mass spectrometry (nanoESI MS) 51. Nanoelectrospray tandem mass spectrometry (nanoESI-MS/MS) 52. Nanospray ionization (NSI) 53. Newborn Screening Ontario (NSO) 54. Outer diameter (OD) 55. Parylene-glass nanospray ionization (PG-NSI) 56. Polycarbonated (PC) 57. Polydimethylsiloxane (PDMS) 58. Polymerase chain reaction (PCR) 59. Polymethyl methacrylate (PMMA) 60. Polytetrafluoroethylene (PTFE, or Teflon) 61. Porous polymer monoliths (PPMs) 62. Post translational modifications (PTMs) 63. Reactive ion etching (RIE) 64. Rotation per minutes (RPM) 65. Scanning electron microscopy (SEM) 66. Signal-to-noise ratio (S/N) 67. Sinapinic acid (SA) 68. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) 69. Specific surface area (SSA) 70. Succinylacetone (SA) 71. Tandem MS (MS/MS) 72. Total ion count (TIC) 73. Trifluoroacetic acid (TFA) 74. Ultraviolet (UV)

1

Chapter 1 Review of Microfluidic Interfaces for Mass Spectrometry

1.1 Mass Spectrometry

In the last decade, mass spectrometry (MS) has become one of the most powerful tools

for bioanalysis. Not only can it provide qualitative information such as the elemental

composition, structure, and isotopic abundances for a given sample, but it is also useful for

quantitative determination of molecular weight and amount of the analyte. The growth in

popularity was driven by the development of two ionization methods useful for producing ions

from nonvolatile biomolecules: electrospray ionization1 (ESI) and matrix assisted laser

desorption ionization (MALDI)2,3. These so-called soft ionization techniques enabled, for the

first time, the capacity to analyze large biomolecules such as proteins (>0.5MDa).4

1.1.1 Ionization Methods

In ESI, an electric field is applied to the tip of a fluid filled tube (i.e., a pulled or

sharpened capillary) to generate an aerosol of charged droplets. In ESI,5 electrical field causes

charge accumulation on the surface of the liquid that is pushed through the nozzle. When the

forces generated by the charge repulsion on the surface overcome those associated with surface

tension, the liquid transforms into a cone shape called a Taylor cone. An aerosol of droplets is

generated at the tip of the cone where the electric field is the strongest. Once released from the

nozzle, each charged droplet begins to shrink as the solvent evaporates. This process is often

accelerated by using a heated chamber and/or a sheath gas flow along the spray trajectory. As

droplet size decreases, molecular ions are pushed closer together. As the columbic repulsion

2

force approaches that of the surface tension, the droplet becomes unstable and disintegrates. This

process repeats itself until the droplet has split to the point where it carries only a single, charged

molecule.5,6

Typically, in MALDI, solutions of analytes are mixed with solutions of a high

concentration of matrix, an additive chosen because of its tendency to form well-defined crystals

and to absorb ultraviolet (UV) light. The mixture is allowed to precipitate and crystallize on a

surface, where it is subsequently analyzed. As the crystal is irradiated with laser light, analyte

and matrix molecules are desorbed off the surface because of heating. There are some competing

ideas about the mechanism, but in one that is commonly cited7, it is believed that the desorbed

matrix molecules also become ionized as a result of the laser energy, and that this energy is then

transferred to the desorbed analyte, which in turn becomes ionized. This process is typically

performed under vacuum and the charged analytes/matrices are guided to the MS through an

electric field for analysis.6

Although ESI and MALDI share some fundamental similarities, each has distinct

advantages and disadvantages such that they are complementary for many applications. For

instance, both offer a rapid and accurate determination of molecular weight for a wide range of

compounds. However, ESI has the advantage of generating multiple charged states; hence, a

more accurate molecular weight can be obtained from the distribution of these multiply charged

peaks.1 Both techniques have the ability to ionize intact multi-molecule complexes (e.g., protein

complexes8); however, in the presence of high concentration salts (often required to stabilize the

structure of protein complexes) or other unwanted constitutents (ie. surfactants for solubilizing

membrane proteins), the formation of ions can be suppressed making the analysis impossible.9,10

3

This effect is less pronounced in MALDI, in which ionized products can be detected even in

presence of salts (e.g., 100mM). Finally, in MALDI, signal intensity is often difficult to correlate

to the amount of analyte in a sample because of “spot-to-spot” variations; this contrasts with ESI,

where signal intensity increases linearly with the analyte concentration over a wide range

(typical dynamic range for quadruple instruments are 4-5 orders of magnitude). Note that the

ionization efficiencies can vary widely as a consequence of experimental conditions for both ESI

and MALDI, making quantitation a challenge for both techniques.

1.1.2 Mass Spectrometry for Proteomics

MS has been used for many decades for small molecule applications including

elucidation of the structure of synthetic organic molecules,11-14 identification of drugs of abuse

and drug metabolites in biological fluids (e.g., blood, urine, and saliva),15,16 respiratory gas

analysis,17-19 dating archaeological specimens,20-23 and determination of atmospheric composition

in space exploration.24,25 However, in the past 2 decades, since the introduction of ESI and

MALDI, there has been an explosive growth in applying MS for high-throughput biomolecule

analysis.

MS has become arguably the most important technique used for proteomics,26-32 the

systematic study of all of the proteins present in a given system (e.g., cells, tissues, organisms).

With the development of new tandem MS (MS/MS) techniques, mass spectrometric

identification of proteins has essentially replaced the classical technique of Edman degradation in

protein chemistry,27 because MS is much more sensitive and can deal with protein mixtures with

much higher throughput. However, MS alone cannot handle the entire proteome, a complex

system that may contain over 20,000 different proteins, not to mention many structurally similar

4

but functionally different isomers formed by post translational modifications (PTMs).28 To

complicate matters, the proteins in a proteome have a wide dynamic concentration range, pI,

polarity, and solubility, making global experimental protocols virtually impossible to develop.28

Hence, MS-based proteomic sample analysis often requires coupling with various separation

techniques such as 2D gel electrophoresis (2DGE)26,30 and 2D liquid chromatography

(2DLC).31,32

In 2DGE-MS analysis, a crude protein mixture (e.g., from cell extract) is applied to a

‘first dimension’ gel strip that separates the proteins based on their isoelectric points. After this

step, the now separated proteins are applied to a ‘second dimension’ – sodium dodecyl sulfate

polyacrylamide gel electrophoresis (SDS–PAGE) gel, where proteins are denatured and

separated on the basis of size. The gels are then fixed and the proteins visualized by silver

staining. The relevant protein bands are excised and subjected to reduction, alkylation, and

digestion, after which the sample is purified and concentrated for analysis using either ESI or

MALDI-MS. These processes are then repeated hundreds of times to analyze each relevant spot

on the gel.

While 2DGE-MS is commonly used, an alternative approach of 2DLC-MS, also referred

to as “shotgun proteomics”, has recently become popular.31,32 In this technique, protein samples

are first chemically processed, purified and concentrated before separating by 2DLC (typically

strong cationic exchange followed by reverse phase). Afterwards, separated peptides are directly

introduced to MS (typically by ESI) for online analysis. A key advantage of this technique is that

it eliminates the tedious process of band selection and excision, a particularly time-consuming

bottleneck for 2DGE based methods. However, the drawback of this method is it requires very

5

complex algorithms (e.g., SEQUEST33,34) to identify proteins from millions of peptides that were

sequenced by MS/MS.

Regardless of which method is used, all proteomic samples require some degree of

cleanup and processing prior to MS analysis. These processes are extremely tedious and time-

consuming; hence, they are attractive targets for automation and integration by microfluidics.35

6

1.2 Microfluidics

Over the past decade, microfluidics has emerged as a new analytical tool for many

chemical and biological applications. The main concept of microfluidics is the ability to

manipulate and control small volumes of fluids at the micrometer scale. In comparison to

conventional macroscale techniques, microfluidics offers many advantages such as reduced

sample and reagent consumption, faster kinetics and thus reduced analysis time, the capability to

integrate multiple functional components, portability of system, and potential for full

automation.36,37 The possibility to integrate different steps of the whole analytical procedure on a

single device results in so-called micro total analytical systems (µTAS), or “lab on a chip”

(LOC).38 Although to date there are limited examples of LOC, microfluidic techniques continue

to gain popularity as alternatives. Currently, there are two paradigms of microfluidics: channel

microfluidics and digital microfluidics.

1.2.1 Channel Microfluidics

In channel microfluidics, fluids are manipulated inside micron-dimension channels. Fluid

flow in these microfluidic devices can be continuous or discontinuous/segmented. Continuous

fluid flow can be achieved by capillary action,39-42 external pressure sources43-46 (e.g., pumps,

syringes, etc.), internal pressure sources47-52 (e.g., microvalves, gas expansion principles, etc.),

and various electrokinetic mechanisms.53-57 Continuous flow has been applied to many lab-on-a-

chip applications including lateral assay tests (LAT),39,40 hydrodynamic focusing for sorting43,44

and counting46,47 micro-scale objects, and high resolution and efficient chemical separations56,57

(by far the most popular application). On the other hand, discontinuous or segmented flow

7

systems are mainly powered through external mechanical pumps. These systems are able to

generate small liquid segments which act as individual microreaction vessels for confinement.

This leads to reduced reagent consumption as well as the ability to perform large number of

different experiments within a short period of time, which makes the platform a promising

candidate for high-throughput screening applications.58-61

The following section presents a brief review of some applications for continuous flow

based microfluidic systems, as well as an introduction to some integrated microchannel

platforms.

1.2.1.1 Applications of Continuous Flow based Microfluidics

With decreasing length scale, surface phenomena (e.g., surface tension, capillary forces,

etc.) become increasingly dominant over volume phenomena (inertia forces, vortices, etc.). This

permits purely passive fluid flow based on capillary action used in popular lateral flow assays

that are known as test strips (e.g., the pregnancy test strip).39 For example, in a typical test strip,

liquid movement is controlled by wettability and the feature size of the porous or microstructured

substrate. The substrate contains pre-loaded chemicals (e.g., antibodies, reactive reagents) that

are necessary to perform the specific chemical or immunological assays on the device. The

readout of an assay is typically done optically and is implemented through a color change in the

detection area that can be seen by the naked eye. A drawback of the LAT format is its simplicity.

Assay protocols within the capillary driven systems follow a fixed process; thus, a limited

number of applications can be adopted for a given design. Moreover, the dependence of sample

movement on purely capillary action can also be a problem, leading to false positive or negative

results, and decreased precision.62 Nonetheless, the fact that 6 billion glucose test strips were

8

sold in 200763 indicates that LAT is a gold-standard microfluidic platform in terms of cost,

handling simplicity, and robustness.

Another effect of moving liquids at micro-length scale is the onset of laminar flow at low

Reynolds numbers - a ratio of inertial forces over viscous forces. Under laminar flow conditions,

fluid mixing is limited to diffusion and this enables the creation of well-defined liquid-liquid

interfaces down to cellular dimensions. This combination of controllable diffusion mixing and

stable phase arrangements has led to the development of hydrodynamic focusing technology, a

technique used to align particles or cells in continuous flow for analysis and sorting in flow

cytometry.45 The downside of the flow focusing technique is Taylor dispersion of samples which

can make it challenging to accurately track analyte concentrations. Although improvements have

been made to minimize dispersion by reducing the channel diameter, narrowing the channel

increases the risk of clogging as well as buildup of back pressure and shear stress.64

Perhaps the foremost popular application for continuous fluid flow in microchannel

systems is chemical separations. High efficiency, high resolution separations can be obtained

using either pressure-driven flow or electroosmotic flow (EOF).65 The former typically requires

complex fittings and valves that may compromise the benefits of using microchannels. In the

latter, voltage-driven EOF is used to pump the mobile phase through an open channel or through

a channel filled with a solid stationary phase in a technique called microchannel

electrochromatography (MEC).66,67 For electrokinetic flow, small channels have the advantage of

a high surface-to-volume ratio, and thus dissipate heat more efficiently than large counterparts.

In addition, the flat velocity profiles of EOF flow enables high efficiency separations with very

little amount of time. For example, Jacobson68 demonstrated an electrophoretic separation of two

9

fluorescent dyes in under 150 ms in a open glass microfluidic channel. Similarly, Ericson et al65

first reported the use of porous polymer monolith (PPM) as a stationary phase for MEC. These

PPMs were formed by thermally initiated free-radical polymerization of acrylate based

monomers yielding a highly macroporous continuous rod. Systems with approximately 500,000

theoretical plates have been reported for systems with these types of PPMs incorporated in

microchannels.69 Even though electrophoretic separation in microchannels benefits from faster

heat dissipation and more efficient separation, technical challenges such as buffer incompatibility

(EOF flow depends on buffer pH and ionic strength), changes in pH as a function of electrolysis,

and changes in EOF velocity as a function of evaporation of solvent are still problematic.

1.2.1.2 Integrations of Channel Microfluidics

As discussed previously, a key feature in LOC is the possibility to integrate multiple

modules on a single platform. Traditionally, this was done through the use of repeated

photolithographic steps (ie. 3D photolithography) to generate a multilayer structure. For instance,

as shown in Figure 1.1a, Xie et al.51 made a microfluidic chip that integrates all fluidic

components of a gradient liquid chromatography system. The fabricated chip included three

electrolysis-based electrochemical pumps50 (one for loading sample, and the other two for

delivering solvent gradient), a sample injector, a mixer, a column packed with silica beads, an

integrated frit for bead capture, an electrospray nozzle, and platinum electrodes for delivering

current to both pumps and generating an electrospray. Although the microchip took several days

with many photolithography steps to fabricate, the system had comparable performance with a

commercial nanoLC system.

10

Recently, two particular developments in microfabrication technologies have accelerated

the generation of integrated micofluidic devices. First is the development of soft lithography,78,79

which enables quicker, less expensive creation of prototypes using elastomeric polymers such as

polydimethylsiloxane (PDMS).70,71 The second development is multilayer soft lithography,72,73

which enables the creation of pneumatic valves, mixers and pumps that can be used to actuate

fluids in microchannels. Such elastomeric actuators have enabled the design of large-scale

integrated microchannel systems for high-throughput processes. For example, Thorsen et al.73

have developed elastomeric PDMS devices with thousands of valves and hundreds of

individually addressable chambers for parallel high throughput screening of bacterial cells

expressing an enzyme of interest (Figure 1.1b). In later studies, they developed similar

mechanical valve-based systems for the automation of cell isolation, cell lysis, DNA purification

and DNA recovery without any sample treatment on a single microfluidic chip.49

Although microchannel systems are a good fit for many applications discussed above,

they are less suitable for tasks that require a high degree of flexibility in fluid manipulation. In

addition, microvalves and related technologies are inherently complex and complicated to use,

which has led many scientists to avoid using this technology. Unfortunately, microvalves are

important; closed-channel systems without valves are inherently difficult to integrate and scale

because the parameters that govern flow field vary along the flow path making the fluid flow at

any one location dependent on the properties of the entire system (e.g., clogging74). Moreover,

many microchannel applications are analytical rather than preparative as samples handled in

microchannel are difficult to recover for further processing. These technical challenges have led

to the development of an alternative paradigm of microfluidics called digital microfluidics.

11

a)

b)

Figure 1.1: Examples of integrated microchannel microfluidics. (a) Photograph of fabricated microfluidics LC chip for proteomic applications. Reproduced from reference 51 with permission. Copyright © 2005 The American Chemistry Society. (b) Microfluidic large-scale integration: picture of a microfluidic chip used for parallelized high-throughput screening of fluorescence-based single-cell assays Reproduced from reference 49 with permission. Copyright © 2007 Nature Publishing Group.

12

1.2.2 Digital Microfluidics

Digital microfluidics (DMF) shares many advantages with microchannel-based fluidics,

such as reduced reagent/sample consumption, fast reaction kinetics, and capacity to integrate

multiple components for lab-on-a-chip applications. Nonetheless, these two techniques are

fundamentally different. First, DMF is an open system, where droplets are manipulated over an

array of patterned electrodes coated with an insulating dielectric.75-78 Although microchannels

have been used to manipulate droplets,59-61,79-82 DMF is distinct in the sense that droplets are

addressed individually rather than in series, which allows for droplets to be dispensed, merged,

mixed and split independently from each other. A second difference is reagent isolation -

droplets serve as discrete microvessels, in which reactions can be carried out without cross-talk

between samples or reagents; this stands in contrast to microchannels, which are prone to

undesirable hydrostatic and capillary flows.83 A third difference is the geometry – as DMF is

inherently an array-based technique, it is a good match for array-based biochemical applications.

Finally, since droplets are manipulated on an array platform, droplets’ paths are reconfigurable to

suit different applications.

There are two general DMF device geometries, closed (Figure 1.2a) and open (Figure

1.2b) formats. In the closed format, also known as two-plate DMF, droplets are sandwiched

between the ground plate (top) and DMF substrate with patterned electrodes (bottom). In the

open format, also known as single-plate DMF, droplets are positioned on a single DMF substrate

which houses both ground and actuation electrodes in a side-by-side geometry. In both

geometries, droplets are isolated from the electrodes by an insulating, dielectric coating. In

addition to the difference in the position of ground electrodes, the open format facilitates faster

13

sample and reagent mixing, and is easily accessible to external detectors; however, it suffers

from increased sample evaporation and inability to perform droplet dispensing and splitting. In

contrast, the closed format can perform all droplet operations (i.e., dispensing, merging, splitting,

etc).

b)a)

Figure 1.2: Digital microfluidics. Schematics of closed (a) and open (b) DMF devices. Reproduced with permission from 84. Copyright 2008 The Royal Society of Chemistry.

DMF devices are typically fabricated using standard photolithography processes and wet

etching techniques in clean room facility. However, efforts have been made to fabricate DMF

devices from inexpensive, accessible methods such as micro-contact printing,85, laser printing,86

and rapid marker masking techniques.87 These techniques have made DMF accessible to

academic and industrial groups that do not have access to clean room facilities, and is

transforming DMF from being a curiosity for aficionados into a technology that is useful for

biochemical applications at large.

1.2.2.1 Applications of Digital Microfluidics

DMF allows for precise spatial and temporal control of fluids, making it amenable to

complex biological reaction schemes such as enzyme assays,88-90 proteomic sample processing.91-

96 cell assays,97-100 immunoassays,101-103 and real time PCR.104 However, a nontrivial challenge in

14

implementing biological reactions on DMF is non-specific adsorption of biological molecules.

This is commonly referred to as surface fouling, and it can lead to cross contamination, sample

loss, and even droplet sticking, which renders DMF devices useless. To overcome this challenge,

Srinivasan et al.88 demonstrated that fouling can be prevented by suspending droplets in an

immiscible oil, which serves as a liquid barrier between droplets and the device surface. More

recently, the Wheeler group has developed two alternative strategies to minimize surface fouling

– (1) the use of droplet additives,105 and (2) the use of removable polymer coverings106 (see

Chapter 5).

Enzyme assays were the first applications implemented on DMF platforms. Taniguchi et

al.89 demonstrated a bioluminescence assay for luciferase in the presence of adenosine

triphosphate. More recently, the Fair group at Duke University demonstrated a fully automated

glucose assay in a range of physiological fluids (serum, saliva, plasma, and urine) on a DMF

device.88 In this study, droplets of glucose oxidase were merged with sample droplets spiked

with glucose, then mixed, and the glucose concentration was measured using an integrated light

emitting diode/photodiode detector (Figure 1.3a). Finally, the Wheeler group applied DMF to

study enzyme kinetics. As proof of principle, separate droplets of alkaline phosphatase and

fluorescein diphosphate were merged and mixed on a multiplexed DMF device (Figure 1.3b).90

Enzyme reaction coefficients, Km and kcat generated by the DMF technique agreed with literature

values. The assays used much smaller volumes, and had higher sensitivity relative to

conventional methods.

For proteomic sample processing, protein samples are typically subjected to extraction

via precipitation, reduction, alkylation, enzymatic digestion, purification, and resolubilization.

15

While a completely integrated proteomic workup by DMF has not yet been implemented, the

field is moving in this direction. For example, Jebrail et al.91 developed a DMF-based protocol

for extracting and purifying proteins from serum by precipitation, rinsing, and resolubilization

(Figure 1.3c). The method had comparable protein recovery efficiencies (≥80 %) relative to

conventional techniques, combined with the advantages of no centrifugation, and faster

extraction and purification. Following this study, Luk et al.92 and Chatterjee et al.96 reported

multi-step protein processing techniques in which protein samples were reduced, alkylated, and

digested sequentially on chip prior to MALDI-MS analysis (Figure 1.3d). In related work, the

Garrell and Kim groups at UCLA93-95 developed DMF-based methods to purify peptides and

proteins from heterogeneous mixtures. The methods comprised a series of steps, including drying

sample droplets onto a device surface, rinsing the dried spot with deionized water droplets to

remove impurities, and finally delivering a droplet of MALDI matrix to the purified proteins for

analysis on-chip. These represented important milestones for DMF-based sample purification,

and led to the development of digital microfluidic solid phase extraction (DMF-SPE) method

described in the Appendix.

16

a)

b)

d)

c)

e)

f)

Figure 1.3: Biological applications of DMF. (a) Picture of DMF device used to perform glucose assays. Reproduced with permission from 88. Copyright 2004 The Royal Society of Chemistry. (b) Picture of a multiplexed DMF device used to study enzyme kinetics. Reproduced with permission from 90. Copyright 2008 The American Chemical Society. (c) Picture of DMF device for protein precipitation in which a droplet of supernatant liquid is driven away from a solid precipitate. Reproduced with permission from 91. Copyright 2009 The American Chemical Society (d) Frames from a movie of a DMF device used to perform multi-step protein processing, in which protein samples were reduced, alkylated, and digested sequentially on chip. Reproduced with permission from 92. Copyright 2009 The American Chemical Society. (e) Pictures of a DMF device used for cell-based toxicity assays. Reproduced with permission from 98. Copyright 2008 The Royal Society of Chemistry. (f) Pictures of a digital microfluidic platform developed for complete cell culture. Reproduced with permission from 97. Copyright © 2010 The Royal Society of Chemistry.

17

Another increasingly popular application for DMF is cell based assays, as the reagents

and cell media materials are often prohibitively expensive for high-throughput techniques. The

first cell based assays on DMF was conducted by Barbulovic-Nad et al.98 In this work, a toxicity

assay was performed on a DMF platform in which droplets containing cultured Jurkat-T cells

were merged with droplets containing different concentrations of Tween-20, a surfactant that is

lethal to cells. A viability dye was subsequently mixed with the droplets (Figure 1.3e) and the

resultant mixture was analyzed using a fluorescence plate reader. The results indicated that DMF

based cell assays were more sensitive than identical assays performed in 384 well plates. In

addition, it was shown that DMF actuation had no significant effect on cell vitality. More

recently, Barbulovic-Nad et al.97 developed the first microfluidic platform capable of

implementing all of the steps required for mammalian cell culture—cell seeding, growth,

detachment, and re-seeding on a fresh surface. This technique demonstrated cell growth

characteristics comparable to those found in conventional tissue culture and were used for on-

chip transfection of cells (Figure 1.3f).

Immunoassays have been another popular target for implementation by DMF. For

example, Sista et al.103 reported the use of DMF to detect insulin and Interleukin-6 using droplets

carrying magnetic beads modified with immobilized antibodies. In this work, droplets containing

magnetic beads bearing antibodies were merged with droplets containing known concentrations

of analyte, and a magnetic field was then used to separate the beads from the supernatant. The

beads were then resuspended in a new buffer droplet, and the immobilized analyte was detected

by chemiluminescence. The assay had low detection limits (0.24 pg µL-1), and standard errors of

less than 3%.103

18

The same group recently developed a DMF platform for multiplexed real time PCR.104

The DMF platform comprises 3 specific regions: a magnetic bead handling region for extraction

of DNA from clinical sample, a thermocycling region in which droplets containing extracted

DNA can be cyclically shuttled in between two fixed temperature zones for amplification, and

dedicated detection spots for real time quantification of amplified DNA using a custom-designed

miniaturized fluorimeter module. Using such system, they were able to detect trace amounts of

DNA of methicillin-resistant Staphylococcus aureus (MRSA), Candida albicans, and

Mycoplasma pneumonia with up to 94.7% amplification efficiency.104 This miniaturized portable

platform has recently been commercialized by Advance Liquid Logic. They speculate this

microfluidic PCR system can be used to detect microbial DNA in clinical specimens for rapid

diagnosis of infectious diseases.107

While the most popular applications of DMF have been in the area of biological

applications, there are growing numbers of non-biological applications that are attracting

attention. For example, Jebrail et al.108 introduced the first two-plate digital microfluidic platform

for chemical synthesis that is suitable for control of many different multi-component, multi-step

reactions in parallel. The platform was used to carry out synchronized synthesis of five peptide

macrocycles from three different components followed by late-stage modification with

thiobenzoic acid to generate aziridine ring-opened products. This technique demonstrated the

potential of using DMF for fast and automated synthesis of libraries of compounds for

applications such as drug discovery. Likewise, Mousa et al. 109 demonstrated the use of DMF for

clinical extraction of estradiol from 1 µL samples of breast tissue homogenate, as well as whole

blood and serum. The method offered up to 4000x reduction in sample size, and 30x faster than

19

conventional extraction method. In addition, this method does not require invasive biopsies of

tissue samples, and suggests that routine monitoring of estrogen level in local breast tissue may

be feasible.

1.2.2.2 Integration in Digital Microfluidics

DMF is a promising technology for applications that involve extensive sample

preparation; however, continued growth in this emerging field depends on advances in system

integration, and its compatibility with existing detection schemes. Since continuous flow

microchannel (which is well suited for analytical applications) and DMF techniques (which is

well suited for preparative applications) are complementary, coupling both techniques would be

a key step toward reaching a goal of fully integrated LOC for inline sample processing and

separation. In response to this challenge, the Wheeler group has made significant strides toward

combining both microfluidic paradigms in a novel "hybrid microfluidics" device

architecture.108,110 Two approaches have been developed, comprising an array of DMF electrodes

interfaced either in a “side-by-side”110 or “top-down”108 configuration with a network of

microchannels. As proof of principle, on chip digestion of model proteins followed by

electrophoretic separation of digested peptides were carried out using both designs. Although

separated peptides were detected using laser induced fluorescence, the results were promising

and the authors anticipated that integration of such hybrid device with MEC and nanoESI-MS

will form a powerful new tool for application to shotgun proteomics and many other

applications. Integrating DMF with a portable capillary electrophoresis analyzer has also been

reported by Gorbatsova111 et al. In this study, DMF acted as an autosampler for transporting

droplets of sample and buffer to a capillary interface for CE separation. The combination of

20

DMF sampling and portable CE analyzer was an important step toward full integration, and this

technique may be useful for high throughput sequencing of nucleic acid libraries.112

Numerous detection schemes including optical, electrochemical detection, and MS (see

section on Microfluidic-MALDI-MS interface for details) have been coupled to DMF processes.

Optical detection is perhaps the most widely employed because it offers a nondestructive

operation mode, capability for multiple sensing, and rapid signal generation and readout. For

example, absorbance detection have been used for immunoassay103 and chemical synthesis,113

and fluorescence and chemiluminescence have been employed for polymerase chain reaction

(PCR)102,114, enzyme assay106, cell-based assay98, and immunoassay.103 However, optical

detection has been limited in sensitivity due to the relatively short optical path lengths in DMF

devices. In addition, electrochemical detection has been integrated with DMF devices. Poulos et

al. combined the DMF droplet actuation with on-chip thin-film Ag/AgCl electrodes for parallel

formation and electrochemical measurement of an artificial lipid bilayer array.115 Unfortunately,

this technique can only detect the electrical properties of analyte species undergoing redox

reaction, and is thus limited to electroactive species. DMF has also been integrated with MS due

to its high sensitivity and the capability to provide exquisite qualitative information when used in

tandem (i.e. MS/MS). Since MS can not be readily miniaturized, much effort has been directed

towards interfacing DMF to MS in an offline mode (See section on Offline Coupling of

Microfluidics and MALDI-MS) (i.e. the analyte is detected either by collecting the sample from

the chip or insert the dissembled chip into the detector instrument). Although there have been

few reports of online coupling (i.e. detects analyte directly on chip), it can be done; in fact,

21

Chapter 3 and 4 in this thesis present two independent methods for coupling DMF with ESI-MS

for direct inline analysis of samples.

22

1.3 Microfluidic Interfaces for Mass Spectrometry

In the past 15 years, there has been great interest in coupling microfluidics with MS.

Because ESI and MALDI are the two widely used ionization methods for analyzing

biomolecules by MS they have been the most popular methods for coupling microfluidic devices

to mass spectrometry. In general, microfluidic chips can be directly coupled to MS via ESI using

pressure driven116-118 or EOF driven119-122 systems to direct the liquid into the instrument.

Interfacing microchips to MALDI is typically performed offline through deposition of the chip

effluent on a MALDI target123 or by ionizing directly from the chip.124 There are a few examples

of in-line MALDI analysis from microfluidic devices through continuous flow125,126 or

mechanical interfaces.127,128 Here, I survey the different kinds of microfluidic-MS interfaces

reported in the literature and discuss the most promising geometries.

1.3.1 Microfluidic-ESI-MS Interfaces

As microfluidic devices handle volumes of liquids on the order of nanoliters, integrating

microchannels with ESI requires dedicated nozzles (sometimes called tips or emitters) capable of

spraying minute volumes. The ideal microfluidic ESI interface should exhibit the following

attributes: (1) high sensitivity over a wide range of analytes; (2) good spray stability; (3) ease of

construction and low cost; (4) minimum dead volume at the interface; (5) durability over the

course of continuous runs; (6) robustness against clogging and solvent/buffer systems; (7)

scalability of the fabrication process. With these goals in mind, a variety of strategies for

integrating microfluidic devices with nanoelectrospray ionization (nanoESI) have been

developed. Figure 1.4 is a schematic diagram depicting various approaches for forming

23

microfluidic-ESI-MS interfaces.129,130 These methods can be broadly classified by how the

electrospray is generated, including: direct spray from channels122,131-138 (Figure 1.4a); spray

from mated, conventional tips116,120,139-143 (Figure 1.4b); and (3) spray from microfabricated tips

(Figure 1.4c).51,117-119,121,144-147

Figure 1.4: Schematic of various microfluidic-ESI-MS interface methods. (a) Spray from chip, (b) Spray from attached ESI-emitter, and (c) Spray from integrated, microfabricated emitter. All reproduced from reference 129 with permission. Copyright © 2009 John Wiley & Sons Inc.

1.3.1.1 Spray from Chip

The simplest approach for interfacing microchannels with mass spectrometry is to

electrospray directly from a channel (i.e., the unmodified edge of a device). Spraying directly

from chip would be ideal, because connections between a microchip and external parts can

increase the dead volume and decrease the efficiency of separations as a consequence. Figure 1.5

shows several examples of these microchips. These types of microchips are typically fabricated

24

using standard photolithography, wet chemical etching, and thermal bonding procedures. An

advantage of this method is the simplicity of fabrication of such devices -- a single

photolithography mask design was sufficient for each of the structures shown in Figure 1.5, and

these methods are typically compatible with batch fabrication.

Although MS emitters formed from the unmodified edge of a device are easy to fabricate,

their spray performances are limited because of eluent spreading at the edge of the chip resulting

from the non-tapered geometry and the hydrophilic nature of the substrate. This limitation can be

overcome by integrating hydrophobic coatings on the edges of the devices.131,134,135 However,

hydrophobic coatings degrade over time; and mechanical cutting of substrates is not compatible

with batch fabrication process. Thus, these challenges have ultimately limited the usefulness of

this technique.

25

a) b)

c) d)

Figure 1.5: Examples of microchip spraying directly from an unmodified edge of a device. (a,b)Pictures of a polycarbonate (PC) chip fabricated by hot embossing from a silicon wafer and astable Taylor cone established at the end of a microchannel in this chip. Reproduced from reference 135 with permission. Copyright © 2004 The Royal Society of Chemistry. (c,d) Schematic of a microchip used to electroosmotically pump fluids and to generate electrospray, and photomicrograph of Taylor cone and electrospray generated at the opening after applying 3kV between the microchip and a target electrode. Reproduced from reference 133 with permission. Copyright © 1997 The American Chemical Society.

1.3.1.2 Spray from a Mated Emitter

A second strategy for interfacing microchannels with mass spectrometry is coupling the

microchannels to conventional pulled glass capillary tips. Because of the tapered geometry of

pulled glass capillary tip, no spreading of fluid at the exit is observed. Another advantage of this

method is the fact that metal coated emitters143 or even stainless steel emitters148 can be used to

simplify the application of high voltage to the device. These devices are capable of efficiently

sampling analytes into the spectrometer, generating mass spectra with sensitivities similar to

26

those of conventional techniques. For example, Lazar et al.139 reported sub-attomole detection of

peptides using a glass microfluidic device mated to a conventional electrospray tip. A major

drawback for this strategy, however, is that dead volumes at the junction of chip and the capillary

emitter compromise the resolution of chemical separations within the microchannel. Moreover,

adhesives are often used to immobilize the capillary onto end of microchannel,121 which can

cause unwanted, contaminating peaks to appear in the mass spectra. As a result, this device

geometry is not likely to be useful for most applications.

1.3.1.3 Spray from Integrated, Microfabricated Emitter

A third strategy for coupling microfluidic devices to ESI-MS is the use of

microfabricated, tapered electrospray tips.51,117-119,121,122,136,138,144-147 These emitters exhibit

similar tip shape to pulled glass capillary to limit fluid spreading at the tip, and they are

fabricated using micromachining processes developed for microelectromechanical systems

(MEMS) technologies.

27

a) b)

c) d)

Figure 1.6: Examples of microfabricated emitters. (a) Cartoon schematic of an Agilent HPLC-Chip II. Reproduced from reference 149 with permission. Copyright © Agilent 2000-2011. (b) Picture of an Advion Biosciences ESI Chip. Reproduced from reference 121 with permission. Copyright © 2000 The American Chemical Society (c) Picture of a micromachined PMMA nozzle. Reproduced from reference 118 with permission. Copyright © 2004 The Royal Society of Chemistry. (d) Picture of a micromachined parylene emitter for ESI MS. Reproduced from reference 147 with permission. Copyright © 2000 The American Chemical Society. Several examples of integrated, microfabricated MS emitters are shown in Figure 1.6,

some of which are available commercially (for example, the device shown in Figure 1.6a from

Agilent Laboratories149 and the device shown in Figure 1.6b from Advion Biosciences121). These

devices are capable of sustaining a stable spray with no dead volume between the channel and

tip. For example, Figure 1.6c shows a micro-milled electrospray nozzle in polymethyl

28

methacrylate (PMMA) fabricated by Schilling et al.118 The performance of the spray is

dependent on the nozzle diameter and apex angle. The best performing tip geometry had a 30 μm

wide nozzle with a 60 angle and was capable of generating a stable spray for several hours.

Licklider et al.147 used vapor-deposited parylene-C to fabricate ESI tips (Figure 1.6d) on silicon

microfluidic devices, enabling integrated liquid chromatography51 with mass spectrometry

detection with comparable performance to conventional techniques. The drawback for the

devices reported by Schilling et al.118 and Licklider et al. 147 is the complexity involved in their

fabrication (i.e., multilayer patterning/developing), requiring many sequential photolithography

steps in a cleanroom.

The most promising microfabricated ESI interface may be the one developed by Yin et

al.,117 which features a nanospray tip formed by laser ablation (355 nm) of a polyimide substrate

(Fig. 1.7a). The fabrication required to form these tips is relatively simple, and the devices

integrate separation and sample enrichment modules which lead to mass spectrometry analysis

with peak resolution, limits of detection and signal-to-noise ratio (S/N) similar to those obtained

by conventional macro-scale methods. Complex protein mixtures from blood plasma were

characterized using this platform, with a detection limit in the low femtomole range; this device

is now commercially available (Agilent Technologies).

1.3.2 Microfluidic-MALDI-MS Interfaces

MALDI is an alternative to ESI for an interface between microfluidic platforms and MS.

The geometry of conventional MALDI detection features arrays of crystallized sample spots on

an open surface, and the process is (in general) performed under vacuum. Thus, MALDI is not

an obvious match for interfacing with microfluidics; despite this, several interfaces with

29

microfluidics have recently been developed. These microchip interfaces can be broken down into

two categories: offline or online coupling. Offline coupling with microchannels can be

accomplished by spotting150-152, spraying,123 centrifuging124, or stamping (specific for DMF

platforms).94,95,153 Online coupling with microfluidics is quite challenging, but can be achieved

using continuous flow125,126, or other mechanical interfaces.127,128

1.3.2.1 Offline Coupling of Microfluidics and MALDI-MS

In offline coupling with MALDI, samples are directly deposited onto a target plate under

atmospheric pressure; the plate is then transferred to a MALDI instrument for analysis under

vacuum. The most commonly used method is spotting, for example, with a robotic target spotter.

Lee et al.150 reported an automated digestion and deposition system formed by mounting a

microfluidic chip inside a commercial MALDI target spotter. The chip contains an immobilized

enzyme microreactor in which proteins were digested and then the products were merged with a

coaxial matrix flow. The resulting mixture was eluted off into an infused stainless steel tube and

spotted onto a MALDI target plate. As a proof of principle, a tryptic digestion of cytochrome c

using the microchip was performed. Peaks from peptide mass mapping revealed a 67% sequence

coverage with an excellent MOWSE (molecular weight search) score.

As an alternative to spotting techniques, electrospray deposition has been demonstrated.

In this method, sample eluent is delivered to a microchip interface123 (similar to the spray from a

mated emitter interface described above) operated at 2-3kV and spray is directed toward the

MALDI target plate positioned at several millimeters away from the spray source. In this

paradigm, matrix solution is added after sample deposition using a custom capillary spotting

tool. The advantages of using a spray source are (1) the small aerosol droplets that arise from

30

spray lead to uniform on-target sample morphology and distribution, and (2) the capacity for

high throughput sample deposition through the use of multiple spray tips.

The methods described above used a conventional MALDI target plate for sample

collection; however, the chip itself can also be used as a MALDI target. For example, a spinning

compact disc (CD) format was developed by Gustafsson et al.124 for microfluidic sample

pretreatment prior to off-line MALDI MS analysis. In this method, chemical reactions and

separations are powered by centrifugal forces on a spinning CD. Fluid flow through the chip can

be controlled by disc rotating speed. As proof-of-principle, the CD technology enabled the

identification of tryptic digests with higher certainty than in identical analyses using

conventional techniques (i. e., steel MALDI target), with better sequence coverage, and the same

resolution and mass accuracy. This platform is now commercially available from Gyros AB.

DMF has been used to process proteomic samples and form arrays of spots for analysis

by MALDI-MS.94,95,153 These spots can be transferred onto MALDI target plates through a

stamping process.99,100,153 For instance, Srinivasan et al.153 implemented such a system by

moving the sample droplet to the stamping site that consisted of a loading hole defined in the

DMF top plate. The droplet was then passively transferred onto the MALDI target positioned

above the top plate. Using this technique, MS spectra of protein calibration solution were

collected and all proteins in the stamping sample were correctly identified. Similarly, Wheeler et

al.94,95 used MALDI-MS to analyze proteomic samples manipulated using DMF actuation. In this

work, both the protein and the MALDI matrix droplets were manipulated on-chip in air medium,

thus allowing protein stamping and co-crystallization directly on the chip surface. In addition to

stamping method, we have developed a much simpler method to integrate DMF with MALDI-

31

MS for offline analysis (see Chapter 5). The strategy involves the use of removable polymer

skins on which biochemical reactions can be performed.106 After each reaction, the skins can be

peeled off of the device and affixed onto the MALDI target for analysis. This technique not only

extends the lifetime of devices, but also eliminates cross-contamination, and bridges the world-

to-chip interface.

1.3.2.2 Online Coupling of Microfluidics and MALDI-MS

Online coupling of the MALDI technique with microfluidic devices is difficult because

both sample deposition and ionization processes usually take place in vacuum. Moreover,

enclosed microchannels are by definition not accessible to laser desorption/ionization, which

requires an open surface from which analytes can be sampled into the spectrometer. Therefore,

integration of microfluidic chips with MALDI for online analysis requires special precautions.

Several strategies,125-128 however, have been adopted to circumvent these challenges. Brivio et

al125,126 developed an integrated system to desorb analytes directed from enclosed channels

through sub-micron pores in the device cover. Likewise, Musyimi et al.127,128 employed a

rotating ball to transfer analytes from polymer microchannels to a MALDI-MS system without

compromising the vacuum required for mass spectrometry. This approach has the advantage of

decoupling the ionization process from the separation step without compromising the overall

system performance.

32

1.4 Conclusion

Two factors favor the use of electrospray ionization for coupling microfluidic devices to

mass spectrometers.130 The first is the geometric similarity between the conventional pulled glass

capillary tips and the nanospray nozzles developed for microdevices discussed in the previous

section; the second is the ease with which chemical separations in microchannels can be

integrated with real-time detection. Thus, we believe that nanospray ionization techniques are the

most useful for the construction of robust interfaces between microfluidics and mass

spectrometry for most applications in the future. However, MALDI will continue to be an

important tool used with LOC devices, given how amenable it is to high-throughput analysis.

Here, this thesis presents work toward coupling microfluidics to ESI-MS and MALDI-

MS. Chapters 2 and 3 describe two independent methods for interfacing microchannel based

devices to ESI-MS. These methods fall under the "spray from unmodified edge of chip" category

(Figure 1.4a). Chapter 4 describes the first coupling of a DMF device directly to ESI-MS, and

this method falls under spray from the "mated emitter" category (Figure 1.4b). Lastly, chapter 5

describes a novel method of coupling DMF to MALDI-MS for offline analysis. As described

herein, we demonstrated that these methods are useful for analyzing proteomic samples and/or

small-molecule analytes. We propose that these strategies will be generally useful for a wide

range of applications in the future.

33

Chapter 2 A Practical Interface for Microfluidics and Nanoelectrospray Ionization Mass Spectrometry

In this chapter, we describe a new method for fabricating channel microfluidic interfaces

for nanoelectrospray ionization mass spectrometry (nanoESI MS). These emitters were formed

from thin microscope glass substrates (~0.3 mm) and an inert polymer, parylene-C. Using a

single photolithography step, the emitters were formed contiguously with microchannels at the

rectangular corners of the glass substrates, such that minimum dead volume at the tip was

observed. Device performance was demonstrated by evaluating diverse analytes, ranging from

synthetic polymers, to peptides, to nucleic acids. By all criteria, for all analytes, performance was

similar to that of conventional emitters (pulled-glass capillaries and the Agilent HPLC ChipTM)

with the advantage of rapid, batch fabrication of identical devices.

34

2.1 Introduction

There is wide-spread interest in coupling microfluidics to mass spectrometry, with the

goal of leveraging the efficiency and integration inherent in microfluidic methods with the

unparalleled qualitative analytical capacity of modern mass spectrometers. This interest is

heightened for applications in proteomics, as the tools currently used in this discipline fall far

short of the performance needed for high-throughput analysis.154,155 Two sample introduction

methods are commonly used in mass spectrometry for proteomics, ESI and matrix assisted

MALDI. Although microfluidic devices have been integrated with MALDI,94,124,125,127 we

contend that ESI, when implemented in “nanospray ionization” (NSI) mode, is the most suitable

geometry for integration with microfluidics. This assertion springs from the obvious similarities

between the conventional technique of interfacing liquid chromatography eluent to a

spectrometer by means of pulled-glass nanospray tips, and the linear geometry of microfluidic

channels.

Here we report the development of a new microfluidic-NSI device fabricated by plasma

etching of parylene-C on a glass substrate. The new devices, which we call parylene-glass

nanospray ionization (PG-NSI) chips, fall into the first category of microfluidic-ESI interfaces –

spray from chip or unmodified edge of a device. Unlike the previous methods in this category132-

134, eluent spreading in the new geometry is significantly limited by using very thin substrates

and by the fabrication of tips ending at the corners of the devices. Additionally, the new method

requires only a single photolithography step, with no cutting or etching of glass substrates. The

devices are thus very straightforward to fabricate and have comparable performance to both

commercial pulled-glass emitters and the Agilent HPLC ChipTM. For these reasons, we believe

35

this method has the potential to become useful for a wide range of researchers developing

microfluidic-NSI methods for proteomic analyses.

36

2.2 Materials and Methods

2.2.1 Reagents and Materials

Unless otherwise indicated, reagents were from Sigma-Aldrich (Oakville, ON).

Parylene-C dimer and Silane A174 were from Specialty Coating Systems (Indianapolis, IN).

Hexamethyldisilazane (HMDS) was from Shin-Etsu MicroSi (Phoenix, AZ). Shipley S1811

photoresist and MF321 developer were from Rohm and Haas (Marlborough, MA), chromium

was from Kurt J. Lesker Canada (Toronto, ON), and CR-4 chromium etchant was from Cyantek

(Fremont, CA). Microscope slides (75 x 25 mm, 1 mm thick) and cover slips (No. 1, 0.15 mm

thick) were from Fisher Scientific Canada (Ottawa, ON). Plastic transparencies printed on an

outsourced printer (4000 DPI) were used as photolithographic masks.

For mass spectrometry, ultramark 1621 (Thermo Fisher Scientific, Waltham, MA) was

diluted in acetonitrile according to the instructions of the manufacturer (1 µM final

concentration). An HPLC standard containing methionine enkephalin, leucine enkephalin and

angiotensin II was diluted in a 1:1 methanol/deionized (DI) water solution with 0.3% acetic acid

in total volume (1 µM final concentration). Angiotensin I and II were also diluted in the same

manner (10 µM and 1 µM final concentration, respectively). 20-mer pre-desalted DNA

oligonucleotide (5′AGCAGAGCGACCTCAATGAT3′) (1 µM final concentration) was

dissolved in ammonium acetate buffer at pH 7.5, and diluted in 1:1 methanol/DI water. Insulin

from bovine pancreas was dissolved in acetic acid according to the instructions of the

manufacturer and diluted in 1:1 methanol/DI water.

37

2.2.2 Device Fabrication

Glass substrates (microscope slides and cover slips) were cleaned in piranha solution (3:1

conc. sulfuric acid, 30% hydrogen peroxide) for 10 min, dried, and dip coated with Silane A174.

After drying, the substrates were coated with a layer of parylene-C (20 µm) using a vapor

deposition instrument (Specialty Coating Systems) followed by coating with a sacrificial layer of

chromium (300 nm) by e-beam evaporation (Edwards Auto 306, Wilmington, MA). After

cleaning (acetone, methanol and DI water), drying, and priming with HMDS, S-1811 was spin-

coated onto the substrates (5000 rpm, 30 s). After soft-baking (2 min, 104 °C), substrates were

photolithographically patterned by exposure to UV radiation (365 nm, 35 mW/cm2, 30 s) using a

Karl-Suss MA6 mask aligner (Garching, Germany). Note that the non-standard conditions (i.e.,

rapid spin-coating velocity and long exposure time) were optimized to minimize the formation

and effects of beads of photoresist on the edges of the substrates.

After developing in MF321 (~30 s), the exposed chromium was etched in CR-4 (~30 s),

and then the exposed parylene was etched by reactive ion etching (RIE, 150 W, 98 sccm O2, 200

mTorr for 15 min) in a Trion Phantom etcher (Clearwater, FL). The remaining chromium was

then etched away and the devices dried. An additional layer of parylene (2 µm) was deposited

onto the substrates to make all surfaces uniformly hydrophobic. A second set of substrates was

also coated with parylene (5 µm), and then bonded to the patterned substrates in a vacuum oven

(200 °C, 48 h) while held together by a vise (~20 MPa).156 With exception of the bonding, the

described steps required ~48 h per batch of 20 devices and were conducted in a class 100

cleanroom at the Emerging Communications Technology Institute (ECTI) in the University of

Toronto.

38

2.2.3 Device Operation

As shown schematically in Figure 2.1a, the assembled devices had a channel leading

from an inlet reservoir to an outlet tip at the corner of the device (see pictures in Figure 2.1b and

2.1c). Typically, the channels were 20 µm deep, with widths of 300 µm tapering to 60 µm at the

tips. The cross sectional area of the tips was ~1200 µm2. NanoPort connections (Upchurch

Scientific, Oak Harbor, WA) were attached to the device reservoirs and connected to a syringe

pump via a fused silica transfer line (360 µm OD, 100 µm ID, Polymicro, Phoenix, AZ). A 1:1

mixture of methanol and DI water was flowed through the channel for 15 min to clear any

impurities or particulates left in the channel and tip during the fabrication. Devices were imaged

using a CCD camera and a Hitachi S-5200 electron microscope (Hitachi High Technologies

America, Pleasanton, CA).

Figure 2.1: PG-NSI chip. (a) Schematic of the fabricated device showing top and bottom plates prior to bonding. (b) Picture of the channel and tip of a device constructed from cover-slips. (c) SEM picture of the tip.

To evaluate nanospray shape, potentials (3-4.5 kV) were applied via a conductive union

(Upchurch Scientific) relative to a metallic ground electrode ~2 mm distant from the tip. Pictures

were collected via a camera positioned horizontally relative to the devices. The volumes of the

Taylor cones were estimated by extrapolating a symmetric cone from the dimensions in the

picture.

39

2.2.4 Mass Spectrometry

The performance of home-built parylene-glass devices was compared to that of

conventional pulled-capillary tips (FS360-50-30-N-20-CT 360 µm outer diameter (OD)

capillary, 30 µm inner diameter (ID) tip, New Objective, Woburn, MA) and commercially

available polyimide microfluidic chips (G4240-61002, 15 µm ID tip, Agilent, Waldbronn,

Germany; this chip is not packed with chromatographic media). Each type of tip was interfaced

with an LTQ Linear Ion Trap Mass Spectrometer (Thermo Scientific, Waltham, MA), and

nanoelectrospray performance (spray stability, total ion count and sensitivity) was evaluated for

several analytes, as described below. Unless otherwise indicated, analytes were flowed at 500

nL/min, with an applied potential of 3.5 kV and capillary temperature of 250 °C. Capillary

voltages (ranging from 22 to 50 V) and other parameters were varied for each experiment to

optimize the observed signal. The spectra presented were obtained by averaging 10-50

acquisitions (at a rate of 1-6 acquisitions/s), and are representative of separate analyses per

experimental condition. The MS/MS analysis of Angiotensin I was performed at 3.3 kV spray

voltage; 49 V capillary voltage; 170 °C capillary temperature; 120 V tube lens voltage and

collision energy equal to 25%.

40

Glass

Chromium

Parylene

Photoresist

expose and develop

etch chromium

O2 plasma

bonding

Figure 2.2: PG-NSI emitter fabrication process. For clarity, the thicknesses of various layers are not drawn to scale.

41

2.3 Results and Discussion

2.3.1 Device Fabrication and Operation

The single-photolithography-step fabrication technique described in Figure 2.2 is much

faster than conventional methods, enabling the production of >20 identical devices in less than

48 h of work. We contrast this to the many elegant but highly complex methods reported

previously, requiring up to three separate photolithography steps or other labor-intensive

procedures to form operable microfluidic-NSI devices51,118,121,147. The requirement of using

multiple photomasks in these traditional MEMS devices slows production considerably, as

alignment of each new mask to the existing device patterns is tedious and time-consuming. The

new method reported here is also unique in that it is likely the first microfluidic-NSI interface

fabrication procedure to use printed transparencies as photomasks, enabling fast and inexpensive

fabrication relative to traditional chromium-on-quartz masks. A third novelty (which also

contributes to the efficiency of the new method) is the use of pre-formed, inexpensive substrates

(microscope cover slips), which obviates the requirement of high-precision wafer dicing to

release/expose the tips.51,147,157

While the new PG-NSI method compares favorably to previously reported

microfabrication procedures, we note that it is slower than forming conventional, pulled-

capillary emitters (which can be formed in minutes). However, integration of conventional

emitters to microfluidic devices116,119,120,143 leads to the presence of dead volumes, which

adversely affects separations. In the new method described here, the nanospray nozzle is formed

contiguously with the microchannels regardless of the complexity of their design. Thus, we

42

assert that the new method represents a significant advance, allowing for rapid interfacing of

devices with a mass spectrometer without the generation of dead volumes.

2.3.2 Device Performance

As described in the Chapter 1 section on microfluidic interfaces for mass spectrometry,

the new method reported here falls into the first category of microfluidic-NSI devices: spray

from the unmodified edges of microfluidic chips. A critical drawback for devices of this type is

the phenomenon of eluent spreading at the opening of the tips. This is a significant problem for

analyses requiring chemical separations, and is caused by the increased surface area (i.e., the

sides and edges of the tip) that is available for wetting. This phenomenon can be minimized on

microfluidic-NSI tips by using complex, multi-step fabrication techniques to form a tip with

reduced outer diameter118, or by coating the emitter with a hydrophobic surface such as a

fluoropolymer.135 Both measures require additional fabrication time, and coatings typically

degrade after a few uses. In contrast, the PG-NSI devices reported here circumvent the wetting

problem by (1) the use of thin substrates (when assembled, the device is ~300 µm thick), (2)

aligning the tip to the corner of the device, and (3) the use of a hydrophobic substrate (rather than

a temporary coating). When taken together, these factors result in a method with significantly

reduced wetting, and small, stable Taylor cones. In short, the new fabrication method is not only

rapid and straightforward, but also is well-suited for electrospray performance.

To evaluate the amount of eluent spreading at emitter tips, we captured images of sprays

of a 1:1 MeOH/DI water solution in the PG-NSI devices formed from microscope cover slips as

well as from microscope slides (the latter have comparable thickness to devices reported

previously133,135). Potentials were applied between the devices and a ground electrode ~2 mm

43

distant from the tip, and were varied until stable Taylor cones were formed. As shown in Figure

2.3, the sprays formed at the two kinds of tips are quite different, producing droplets of

approximately 2 nL volume for the thin substrates and at least 20 nL for the thick devices. The

former volume is compatible with the peak volumes for microchip separations, making the thin

devices well suited for this application. In addition, when varying the applied voltage and

distance, we observed thicker devices to be more prone to corona discharges,158 leading to

wetting and spray instability. It is clear that the thin devices (i.e., formed from microscope cover

slips) are characterized by superior spray (see Fig. 2.3a) and negligible dead volumes, and thus

these devices were used for the remainder of the work presented here.

(a) Thin substrate (b) Thick substrate

Figure 2.3: Taylor cones (indicated by arrows) produced by devices formed from cover slips (a) and glass slides (b). The droplets forming the spray on the two devices were ~2 nL and ~20 nL, respectively. The reduced area for wetting in (a) contributes to the formation of a small and stable Taylor cone with minimal wetting at the tip.

To characterize the stability of sprays formed at the edge of PG-NSI devices, we

interfaced them to a Thermo LTQ mass spectrometer and evaluated the total ion count (TIC). As

shown in Figure 2.4, the sprays generated by the PG-NSI devices were observed to be stable,

with TICs comparable to those of an Agilent polyimide tip and a pulled glass capillary. Several

PG-NSI devices were evaluated in the course of multiple experiments – typically, they had

44

lifetimes of ~1 week of intermittent use (similar to a pulled-glass capillary), after which they

could be re-coated with 2-5 µm of parylene (by vapor deposition) and used again.

Figures 2.4: Typical total ion current (TIC) traces for a parylene-glass device (a), an Agilent chip (b), and a pulled-capillary glass emitter (c). In each case, the infused solution was 50/50 methanol/DI water with 0.3 % acetic acid. The applied potentials and flow rates for (a-c) were 3.5 kV/0.5 µL/min, 1.4 kV/0.4 µL/min and 2.5 kV/0.6 µL/min, respectively.

45

2.3.3 Mass Spectrometry Performance

The capacity of the PG-NSI devices to infuse samples into the MS for analysis was

evaluated for several analytes ranging from synthetic polymers to peptides and nucleic acids. A

spectrum of calibration standard, ultramark 1621, is shown in Figure 2.5a. Ultramark, a mixture

of fluorinated phosphazenes, is characterized by a series of intense singly charged peaks equally

spaced by m/z 100 and is thus a particularly good benchmark for MS emitters. A spectrum of

several peptides used as HPLC standards, angiotensin II (m/z 1046), leucine enkephalin (m/z

556), and methionine enkephalin (m/z 574), is shown in Figure 2.5b. Singly charged angiotensin

II was also observed, shown in Figure 2.5c. A spectrum of a 20-mer DNA oligonucleotide is

shown in Figure 2.5d – this is of note because analytes carrying large numbers of negative

charge in solution phase (such as DNA) are not typically analyzed by positive mode mass

spectrometry (and thus are typically only observed when using well optimized NSI-MS systems).

In all cases, the analytes could be identified with excellent signal-to-noise ratio.

46

Figure 2.5: Mass spectra generated using PG-NSI devices. (a) Ultramark calibration standard; (b) HPLC peptide standards, including singly protonated leucine enkephalin (556 m/z) and methionine enkephalin (574 m/z), and doubly protonated angiotensin II (524 m/z); (c) Singly protonated angiotensin II (1046 m/z); (d) 20-mer DNA oligonucleotide, [M+5H]5+ (1228 m/z) and [M+4H]4+ (1534 m/z).

In the past decade, tandem mass spectrometry combined with collision-induced

dissociation for peptide sequencing has become a method of choice for proteome profiling. To

demonstrate the compatibility of the new method with peptide sequencing, a PG-NSI device was

used to analyze angiotensin I by MS/MS. As shown in Figure 2.6a, multiple parent ions were

observed in the first mass selection, and after isolation of the triply protonated ion and collision-

47

induced dissociation, several b- and y-ion peptide fragments were identified in the second mass

selection, as shown in Figure 2.6b.

Figure 2.6: Tandem MS spectra generated using PG-NSI devices. (a) Spectra of angiotensin I (1296 m/z) (10 µM) obtained using a parylene-glass device, emphasizing the doubly (649 m/z) and triply (433 m/z) protonated parent ion peaks. (b) MS/MS performed on the 433 m/z parent ion. Several peaks were identified as b- and y-ion peptides (see inset on top right).

Finally, to characterize the detection limits of the new device, we analyzed standard

solutions of insulin. As shown in Figure 2.7, a PG-NSI device can detect 100 and 10 nM

concentrations, with a detection limit of <10 nM. This is comparable to the detection limits of

pulled-capillary and polyimide microfluidic emitter devices (data not shown). In short, the new

PG-NSI devices are similar in all respects to conventional techniques, with the advantage of

rapid, batch fabrication of emitter tips with identical geometries.

48

Figure 2.7: Evaluation of parylene-glass devices for the detection limit of insulin. The peaks at 1148 m/z ([M+5H]5+) and 1434 m/z ([M+4H]4+) are observed with good signal-to-noise ratio at both 100 nM (a) and 10 nM (b) concentrations. The spectrum shown in (b) is an average of only 2 acquisitions.

49

2.4 Conclusion

This chapter presented a new method for fabricating nanospray ionization tips for

interfacing microfluidic devices to mass spectrometers. The construction of these tips is

relatively simple, as the spray is generated from the unmodified edge of a device, not requiring

dicing of substrates to release/expose the tips. However, in contrast to the previous methods of

this type that have been reported, eluent spreading in the new geometry is significantly limited

by (1) the use of thin substrates (when assembled, the device is ~300 µm thick), (2) aligning the

tip to the corner of the device, and (3) the use of a hydrophobic substrate (rather than a

temporary coating). When analytes are sprayed from the new tips, small Taylor cone volumes are

observed, which makes the method attractive for the integration with microfluidic separations.

The devices were evaluated for several analytes, within a concentration range of 10 nM to 10

µM. In all cases, the spectra were comparable to those collected using conventional pulled-glass

capillaries and the Agilent HPLC ChipTM. In on-going work, we are building devices with more

complex channel geometries, to effect sample injections and separations integrated with the NSI

emitters described here, aiming at lab-on-a-chip applications for proteomics analyses.

50

Chapter 3 A Digital Microfluidic Method for Amino Acid Quantification in Dried Blood Spots

Blood samples stored as dried blood spots (DBSs) have emerged as a useful sampling and

storage vehicle for clinical and pharmaceutical analysis in a wide range of applications. For

example, the Newborn Screening Ontario (NSO) facility at the Children's Hospital of Eastern

Ontario evaluates DBS samples from approximately 140,000 babies each year for 28 inherited

diseases. In each screening test, a DBS sample is collected and then mailed to NSO for analysis

by tandem mass spectrometry (MS/MS). Unfortunately, this technique is slowed by an extensive

sample preparation regimen (including excision/punching, extraction, evaporation,

resolubilization, and derivatization), and in addition, high-throughput screening requires robotic

sample handling. As a first-step towards overcoming these challenges, we have developed an

integrated microfluidic system for quantification of amino acids in dried blood spots. The new

method is fast, robust, precise, and is capable of quantifying analytes associated with common

metabolic disorders such as homocystinuria, phenylketonuria, and tyrosinemia. We propose that

the new method can potentially contribute to a new generation of analytical techniques for DBS

samples for newborn screening and other applications.

51

3.1 Introduction

As reviewed in a recent cover story in Chemical and Engineering News,159 blood samples

stored as DBSs have emerged as a useful sampling and storage vehicle for clinical and

pharmaceutical analysis in a wide range of applications. For example, a DBS from every baby

born in the province of Ontario, Canada is analyzed at the NSO facility at the Children’s

Hospital of Eastern Ontario160 for markers for 28 diseases including homocystinuria (build-up of

excess methionine and homocysteine), phenylketonuria (build-up of excess phenylalanine), and

tyrosinemia (build-up of excess tyrosine and its metabolites).161-164 This represents a significant

public health undertaking, requiring the evaluation of approximately 2,800 DBS samples each

week. Many jurisdictions around the world have instituted similar programs.165-167

In each newborn blood spot analysis,160,168-170 a sample is obtained by pricking the

subject’s heel (or by venipuncture) and allowing a spot of blood to dry on filter paper. The DBS

is couriered to NSO, where 3.2 mm diameter circular discs are punched, and the analytes are

extracted, mixed with isotope-labelled internal standards, derivatized, and then reconstituted for

analysis by tandem mass spectrometry (MS/MS). As shown in Figure 3.1a, the derivatization

step transforms each amino acid (AA) to its corresponding butyl ester (derivatized AA) that

allows for a characteristic fragmentation pattern (neutral loss of 102) via collision induced

dissociation (CID). Figure 3.1b-c contains representative primary (MS1) and secondary (MS2)

mass spectra for the amino acid, phenylalanine, with peaks at m/z 222 and 120. In addition to

amino acids, the same derivatization step butylates acylcarnitines (AC), which serve as markers

of inborn errors of fatty acid and organic acid metabolism.164,170

52

(a)

Inte

nsit

y (a

rb)

20016012080m/z

[Phe + H - 102]+ (120)

(b)

(c)

3N HCl in n-butanol

H2N COOH

R

+H3NO

R

O+H2N H

RCID

amino acid (AA) DerivatizedAA Daughter ion

MS1

MS2

Inte

nsity

(ar

b)

250240230220210200m/z

[PheD5 + H]+ (227)

[Phe + H]+ (222)

Figure 3.1: Processing blood samples for quantification of AAs by MS/MS. (a) Reaction scheme involving derivatization of the extracted AA, followed by derivatization with n-butanol, followed by the formation of a daughter ion by CID in the mass spectrometer. (b) Mass spectrum generated from primary analysis (MS1) of derivatized phenyalanine (Phe). (c) Mass spectrum generated from the secondary analysis (MS2) of derivatized Phe showing the loss of 102 amu as a result of CID. The success of DBS and MS/MS for newborn screening has led to a surge in popularity

for similar techniques for a wide spectrum of applications in clinical labs and the pharmaceutical

industry.159 DBS methods allow for the collection of tiny amounts of sample and are convenient

for long-term storage and cataloguing. MS/MS methods allow for the unambiguous identification

and quantification of many different analytes in a single shot. But despite these advantages, a

number of important challenges remain. For example, maintenance of instruments (sample

preparation robots and mass spectrometers) and plumbing (capillary tubes and associated

connections) requires expertise and time.170,171 In addition, the costs (~$20 per sample for

53

newborn screening172) are magnified by the scale of operation (e.g., nearly 140,000 samples/year

for NSO). Finally, collection procedures (i.e., spotting blood samples on filter paper) can suffer

from challenges (e.g., uneven distribution or overlapping blood spots and supersaturated filter

paper) that can skew results.4,160,168 Thus, programs relying on DBS and MS/MS are

continuously seeking innovative laboratory approaches and technologies that can improve

sample preparation throughput and quality of analysis.

Here we report a new automated method for quantifying analytes related to genetic

diseases in newborn blood samples powered by DMF. DMF is a relatively new technique in

which droplets are manipulated on an array of electrodes by application of electromechanical

forces173 and is useful for precise control of complex samples and reagents for biochemical

analyses.174 Moreover, as described previously,175 DMF is particularly useful for extracting

analytes from solid samples, as the solids stick to the device surface, allowing the extracted

analytes to be ferried away for analysis in moving droplets. We report here a series of new DMF-

driven methods for analyzing amino acids in newborn blood samples. As far as we are aware,

this is the first report of microfluidic methods that are capable of direct analysis from DBS

samples. The new methods are automated and have significant advantages relative to the

conventional techniques in terms of sample preparation and reagent use. Moreover, the new

technology is liberated from reliance on robots and capillary-plumbing. We propose that the

methods may someday be a useful tool for newborn screening and other applications.

54

3.2 Materials and Methods

3.2.1 Study Subjects

Liquid blood samples were collected from a healthy adult male volunteer after a 10 h

fasting period, and were kept at –20°C until analysis. Punches from residual dried blood spots

from infants screened by NSO were stored at -80°C until analysis.

3.2.2 Reagents and Materials

L-Methionine (Met), L-Phenylalanine (Phe), L-Tyrosine (Tyr), acetonitrile (ACN),

acetone, methanol (MeOH), boric acid and fluorescamine were purchased from Sigma Chemical

(Oakville, ON). Deuterated Methionine (Met-d3), Phenylalanine (Phe-d5), and Tyrosine (Tyr-

d4) were obtained from Cambridge Isotope Laboratories (Andover, MA). Concentrated

hydrochloric acid (HCl) was purchased from Fisher Scientific (Ottawa, ON) and n-butanol from

ACP Chemicals (Montreal, QC). In all experiments, organic solvents were HPLC grade and

deionized (DI) water had a resistivity of 18 MΩ•cm at 25 °C.

Working solutions of all AAs (25, 50, 100 and 500 µM ea.) were prepared in DI water.

For derivatization of extracted AAs, a 3 N HCl in butanol solution was used. For analysis of AAs

in blood samples, the extracting solvent (MeOH) contained 50 µM of the appropriate deuterated

AA (d3-Met, d5-Phe or d4-Tyr). For quantitative analysis of AA recovery from blood and for

experiments mimicking diseased/healthy infant blood, samples were spiked with 200 µM of the

appropriate AA (Met, Phe or Tyr). A series of 3.2 mm dia. punches of dried blood samples

containing different concentrations of phenylalanine (see Table 3.3) were prepared by NSO

staff.169

55

3.2.3 Reagents and Materials for Device Fabrication

Concentrated sulfuric acid and hydrogen peroxide (30%) were from Fisher Scientific

(Ottawa, ON), Parylene C dimer was from Specialty Coating Systems (Indianapolis, IN) and

Teflon-AF was from DuPont (Wilmington, DE). Shipley S1811 photoresist and MF321

developer were from Rohm and Haas (Marlborough, MA), AZ300T photoresist stripper was

from AZ Electronic Materials (Somerville, NJ), solid chromium was from Kurt J. Lesker Canada

(Toronto, ON), CR-4 chromium etchant was from Cyantek (Fremont, CA), HMDS was from

Shin-Etsu MicroSi (Phoenix, AZ), and Fluorinert was FC-40 from Sigma (Oakville, ON).

Piranha solution was prepared as a 3/1 v/v mixture of sulfuric acid/hydrogen peroxide. Dicing

tape (medium tack) was purchased from Semiconductor Equipment Corp. (Moorpark, CA).

3.2.4 DMF Device Fabrication and Operation

Digital microfluidic devices were fabricated in the University of Toronto ECTI

cleanroom facility, using a transparent photomask printed at Norwood Graphics (Toronto, ON).

Glass devices bearing patterned chromium electrodes were formed by photolithography and

etching as described previously176, and were coated with 2.5 µm of Parylene-C and 50 nm of

Teflon-AF. Parylene-C was applied using a vapor deposition instrument (Specialty Coating

Systems), and Teflon-AF was spin-coated (1% wt/wt in Fluorinert FC-40, 2000 rpm, 60 s)

followed by post-baking on a hot-plate (160 ºC, 10 min). The polymer coatings were removed

from contact pads by gentle scraping with a scalpel to facilitate electrical contact for droplet

actuation. In addition to patterned devices, unpatterned indium tin oxide (ITO) coated glass

substrates (Delta Technologies Ltd, Stillwater, MN) were coated with Teflon-AF (50 nm, as

above).

56

The device design used in methods 1 and 2 (Fig 3.2a,b) featured an array of eighty-eight

actuation electrodes (2.2 × 2.2 mm ea.) connected to ten reservoir electrodes (5 × 5 mm ea.),

with inter-electrode gaps of 40 µm. For method 1, devices were assembled with an unpatterned

ITO–glass top plate and a patterned bottom plate separated by a spacer formed from four pieces

of double-sided tape (total spacer thickness 360 µm). For method 2, the two plates were

separated by six pieces of double-sided tape (total spacer thickness 540 µm). To actuate droplets,

driving potentials (70–100 VRMS) were generated by amplifying the output of a function

generator (Agilent Technologies, Santa Clara, CA) operating at 18 kHz. As described

elsewhere176,177, droplets were sandwiched between the two plates and actuated by applying

driving potentials between the top electrode (ground) and sequential electrodes on the bottom

plate via the exposed contact pads. Droplet actuation was monitored and recorded by a CCD

camera mounted on a lens.

57

Figure 3.2: Three digital microfluidic methods designed to quantify amino acids in dried blood spot samples. In method 1 (a), a 5 µL droplet of blood is spotted directly onto the device surface and allowed to dry. In method 2 (b), a 3.2-mm diameter punch from filter paper bearing dried blood is positioned on the device surface. In method 3 (c), a microchannel nanoelectrospray emitter is coupled to the device to facilitate inline analysis by MS/MS.

58

3.2.5 Hybrid Device Fabrication and Operation

Hybrid DMF-microchannel devices used for method 3 (Fig. 3.2c), were fabricated in four

steps. First, a DMF bottom substrate (layer 2 in Figure 3.3) was fabricated as described above

(but with no Teflon-AF coating). The design, shown in Figure 3.3, was similar to that of the

DMF-only devices, but with fewer electrodes – 2 rows of 9 actuation electrodes (2.2 x 2.2 mm)

and 3 reservoir electrodes (5 x 5 mm). Moreover, the substrates were first modified by drilling an

access hole (~2 mm diameter) through the substrate using a micro drill-press before the

photolithographic processes. After patterning the electrodes, the opposite side was first coated

with 7µm of Parylene-C for bonding with a second substrate (layer 3 in Figure 3.3). Second, a

glass substrate bearing a microchannel nanoelectrospray tip (layer 3 in Figure 3.3) formed in

Parylene was fabricated using methods similar to those described previously (See Chapter 2), but

with a few modifications. 37 grams of Parylene-C were deposited on piranha cleaned, silanized

glass slide (25 x 55 mm) via vapour deposition. After Cr deposition, a microchannel (25 μm wide

x 5 mm long) was photolithographically patterned on the substrate by UV radiation (365 nm, 35

mW/cm2, 50s) using a Karl-Suss MA6 mask aligner (Garching, Germany). Third, the channel

side of layer 3 was mated to the non-electrode side of layer 2, placed under pressure in a

precision vise (~20 MPa), and thermally bonded in a vacuum oven (200 ˚C, 24 h). After cooling,

the top of layer 2 was first coated with 2 μm of parylene followed by spin-coating 50 nm of

Teflon-AF with a small piece of dicing tape covering the accessing hole. The tape was removed

before post-baking on a hot plate (160 ˚C, 10 min). Fourth, the top plate (layer 1 in Figure 3.3)

was assembled with spacers formed from four pieces of double-sided tape as described above for

droplet actuation.

59

Layer 1: DMF Counter Electrode

Layer 2: DMF Driving Electrodes

Layer 3: Microchannel

Access Hole

Spacer

Figure 3.3: Exploded view schematic of the hybrid DMF-microchannel device used for method 3 for in-line analysis by MS.

3.2.6 Non-DMF Sample Processing and Analysis

AAs were extracted and quantified from dried blood spot according to the routine

newborn screening method.169

3.2.7 DMF-Driven Sample Processing

For samples analyzed by DMF methods 1 and 3 (Fig. 3.2a,c), 5-µL droplets of blood

were spotted onto the bottom plate of a device and dried. The top plate was then affixed and two

solvents were loaded into the appropriate reservoirs, including MeOH containing 50 µM of

deuterated AAs (extraction solvent), and 3 N HCl-butanol (derivatization reagent). A reservoir

volume (10 µL) of extraction solvent was dispensed and driven by DMF to the dried sample and

allowed to incubate (5 min). The extraction solvent was then actuated away from the sample and

evaporated to dryness (~15 min, room temperature) at a second site, after which a reservoir

volume (10 µL) of derivatization solvent was dispensed to the dried extract and incubated for 15

min at 75ºC. Following the reaction, the top plate was removed and the solvent was allowed to

evaporate (~15 min, room temperature). For samples analyzed by method 2 (Fig. 3.2b), the

process was identical to the above, but with two differences: (1) punches (3.2 mm dia.) from

60

filter paper bearing dried blood were positioned on the bottom plate of a device in place of liquid

blood, and (2) a larger volume (15 µL) of extraction solvent was used.

3.2.8 DMF-Driven Sample Analysis

Samples processed by digital microfluidic methods 1 and 2 (Fig. 3.2a,b) were analyzed

offline by nanoelectrospray tandem mass spectrometry (nanoESI-MS/MS). Such samples (stored

dry on device or in centrifuge tube until analysis) were reconstituted in 70 µL of

acetonitrile/water (4:1 v/v); samples originating from blood were, in addition, passed through

PVDF membrane centrifuge-filters with 0.1 µm pore diameter (Millipore, ON). Samples were

injected into an LTQ Mass Spectrometer (Thermo Scientific) via a fused silica capillary transfer

line (100 μm ID) mated to a New Objective Inc. (Woburn, MA) nanoelectrospray emitter (100

μm ID tapering to 30 μm ID.) at a flow rate of 0.8 μL min-1, with an applied voltage of 1.7-1.9

kV and capillary temperature of 200°C. MS/MS analysis was carried out by introducing 30%

collision energy to the parent ions and then the fragments over the m/z range of 100-300 were

scanned. AA daughter ions detected in the second mass selection, which exhibit a loss of

butylformate (HCOOC4H9, 102 m/z), were observed and used for quantification. Spectra were

collected as an average of 50 acquisitions, and replicate spectra were obtained for DMF-

derivatized samples of both control and blood. For samples processed by method 3 (Fig. 3.2c),

devices bearing an integrated nESI emitter were mounted on a 3-axis micromanipulator (Edmund

Optics, NJ) positioned near the inlet of the LTQ MS. After sample processing, a spray was

generated by applying 2.5-3.0 kV to a platinum wire inserted in the access hole, with parameters

identical to the above.

61

For samples processed by DMF method 1 (Fig. 3.2a), calibration plots were generated by

plotting the MS/MS intensity ratio of daughter ions from the extracted AAs relative to the those

of the internal standards (i.e., Met m/z 104:107, Phe m/z 120:125, and Tyr m/z 136:140) as a

function of AA concentration in standard solutions (25-500 µM in DI water). Data points

included in the calibration plots represent an average of at least 4 replicate measurements, and

the data in each plot were fit with a linear regression. Blood samples were then evaluated (with

on-chip derivatization and extraction, and measurement by MS/MS relative to internal standards,

as above), and the values were compared to the calibration plots to determine the AA

concentrations.

For samples processed by DMF method 2 (Fig. 3.2b), quantification was similar to that

for method 1, except that standards were formed from adult male blood spiked with Phe to

different concentrations (25-900 µM). 3.1 µL aliquots were pipetted onto pre-punched (3.2 mm

diameter) discs of Whatman 903 filter paper (Whatman, NJ) and were allowed to dry.

Calibration plots were formed with the assumption that all samples contained 38 µM Phe (as per

Table 3.1). Newborn blood samples (as DBS filter paper punches) from NSO were then analyzed

as above and compared with the calibration plot to determine the Phe concentrations.

3.2.9 DMF-Driven Sample Recovery Evaluation

The % recovery of AAs in Method 1 (Fig. 3.2a) was evaluated quantitatively using (i) a

fluorescence-based assay and (ii) MS/MS. For (i), control samples (Met, Phe or Tyr; 50 µM of

each) were processed by DMF (as above), excluding the derivatization step. The dried extracts

were diluted into 95 µL aliquots of borate buffer (20 mM, pH 8.5) in wells in a 96-well

microplate. Upon addition of 5 μL of fluorescamine (5 mg/mL in acetone) the microplate was

62

inserted into a fluorescence microplate reader (Pherastar, BMG Labtech, Durham, NC) equipped

with a module for 390 nm excitation and 510 nm emission. The plate was shaken (5 s) and the

fluorescence was measured. As a control, identical samples that had not been extracted were

evaluated using the same fluorescent assay. To ensure that controls were processed in identical

manner relative to extracted samples, each control sample was spotted on a device, dried and

then reconstituted in buffer for analyses. Four replicate measurements were made for each

sample and control. For (ii), blood samples of known AA concentrations were spiked with 200

µM of AA standards and extracted (as above). Knowing the total concentration of AAs in blood

spots (e.g. native methionine concentration plus spiked methionine), % recovery was obtained by

comparing the concentration values (obtained from calibration curves) vs. the known values.

Table 3.1: Concentrations of amino acids in blood of an adulte male volunteer measured by digital microfluidic method 1 (left column) and normal concentration ranges (right column).

Amino Acid Measured Blood

Concentration (µM)

± 1 S.D.

Normal Blood

Concentration (µM)

Methionine 25 ± 2 16-33178,179

Phenylalanine 38 ± 2 41-68178,180

Tyrosine 46 ± 5 45-74178,181

63

3.3 Results

Digital microfluidic methods were developed to automate the quantification of

methionine as a marker for homocystinuria, phenylalanine as a marker for phenylketonuria, and

tyrosine as a marker for tyrosinemia in dried blood samples. (Note that for Type 1 Tyrosinemia,

succinylacetone164 is typically measured as a primary screen.) To accommodate different

sampling and analysis needs, three related methods were developed. As shown in Figure 3.2, in

method 1 and 3, a sample of liquid blood is spotted directly onto a device and allowed to dry. In

method 2, a punched sample of filter paper bearing dried blood is deposited onto a device for

analysis. In methods 1 and 2, analysis is carried out offline, while in method 3, an integrated

nanoelectrospray emitter facilitates inline analysis with mass spectrometry.

64

3.3.1 DMF-Driven Method 1

1 2 3

4 5 6

Derivatization Solvent

Blood Sample

ExtractionSolvent

ExtractionSolvent

Extract Droplet

Extract

Derivatization Solvent

Derivatized Extract

250

200

150

100

50

0Met Phe Tyr

67 µM

120 µM150 µM

Con

cent

ratio

n (µ

M) Normal Spiked

100 µM

(a)

(b)

Figure 3.4: Analysis of AAs in DBS by DMF-driven method 1. (a) Sequence of frames from a movie depicting several stages in sample processing by DMF including: (1) a dried blood sample prior to processing; (2) mixing and incubating an extractant droplet with the sample; (3) a droplet containing sample extract after translation away from the dried sample; (4) a dried extract; (5) mixing and incubating a derivatization reagent droplet with the dried extract; and (6) the dried, derivatized product. (b) Comparison of Met, Phe, and Tyr concentrations in normal (green) and spiked (red) blood samples as biomarkers for homocystinuria, phenylketonuria, and tyrosinemia, respectively. The dashed lines indicate the upper levels for normal concentrations in newborn blood samples. Each data point represents at least four replicate measurements, and error bars represent ±1 S.D.

While the most common method for DBS sample analysis relies on blood dried onto filter

paper, we propose that for some cases, it may be useful to analyze blood samples that are dried

65

directly onto the surface of a device, as in method 1 (Fig. 3.2a). An experiment using method 1 is

depicted in Figure 3.4a. As shown, a blood sample is spotted onto the device, dried, extracted

into methanol containing isotope-labeled standards, and the solvent is allowed to evaporate. The

extract and standards are then derivatized, and the products are isolated by allowing the solvent

to evaporate. The entire process requires 50 min to complete. Samples processed by method 1

were collected and analyzed off-line by nanoESI-MS/MS to quantify AAs. Calibration curves

with R2 greater than 0.996 (See Figure 3.5) were generated for Met, Phe, and Tyr by analyzing

standards processed by DMF at known concentrations from the abundance ratio of each AA to

its deuterated standard peak in the secondary (MS2) spectra. As listed in Table 3.1, when method

1 was used to evaluate blood collected from an adult male volunteer, the values obtained were in

the expected physiological range and the precision in the method was high with coefficients of

variation (CVs) ranging from 5 to 11%.

66

(a)

Concentration (μM)

Rat

io o

f In

tens

ities

(AA

/IS

)5004003002001000

y = 0.0374x - 0.587

R2

= 0.9959

Methionine

5004003002001000

y = 0.032x - 0.4659

R2

= 0.9992

(b)

Concentration (μM)

Phenylalanine

Rat

io o

f In

tens

ities

(AA

/IS

)

5004003002001000

y = 0.025x - 0.1541

R2

= 0.9996

(c)

Concentration (μM)

Tyrosine

Rat

io o

f In

tens

ities

(AA

/IS

)

Figure 3.5: Calibration curves generated by DMF-driven method 1 for quantification of (a) methionine (Met), (b) phenylalanine (Phe), and (c) tyrosine (Tyr). Data were generated by plotting the intensity ratios of the daughter ions of each amino acid (AA) relative to their deuterated internal standard (IS) (i.e., d3-Met, d5-Phe, d4-Tyr, respectively) as a function of AA concentration. Each data point represents at least four replicate measurements, and error bars represent ±1 S.D. Regression lines were linear with R2 > 0.996 for each analyte.

To evaluate the potential of DMF-driven method 1 as a platform for discriminating

between disease and healthy states in blood, spiked blood samples (mimicking diseased states)

67

and non-spiked blood samples (mimicking healthy state) were analyzed by MS. Figure 3.4b

shows a comparison of measured concentration of AAs in normal and spiked blood samples. The

dashed line indicates typical threshold values that correlate with homocysteinuria (100 μM

Met)160,182, phenylketonuria (120 μM Phe)183,184, and tyrosinemia (150 μM Tyr).164,182 As shown,

the method is useful for distinguishing between these concentrations.

Extraction efficiency is an important parameter for any new analytical method. To

evaluate the extraction efficiency of DMF method 1, two orthogonal tests were used:

fluorescence and MS/MS. In the former test, fluorescamine, a fluorogenic reagent that exhibits

no fluorescence until it reacts with primary amines185 was used to label standard samples before

and after extraction to determine concentrations. In the latter test, blood samples were spiked

with AA and recovery was determined by comparing the AA concentration (endogenous plus

spiked AA) vs. known concentration. As listed in Table 3.2, the data from the two orthogonal

methods (fluorescence and MS/MS) agree and reveal the new DMF technique to be very

efficient. Recovery was ≥80% for each sample evaluated. As above, the precision of these

measurements was high, with CVs ranging from 1 to 10%.

68

Table 3.2: % Recovery of DMF-driven method 1 measured by fluorescence (left) and MS/MS (right)

Amino Acid % Recovery by

Fluorescence ± 1 S.D.

% Recovery by MS/MS ± 1

S.D.

Methionine 98 ± 10 100 ± 1

Phenylalanine 86 ± 9 85 ± 5

Tyrosine 82 ± 10 84 ± 7

3.3.2 DMF-Driven Method 2

DMF-driven method 2 (Figure 3.2b) was developed to analyze punched discs from DBS

formed on filter paper. A portion of an experiment is depicted in Figure 3.6a. As shown, a

droplet of extraction solvent was dispensed and driven to a 3.2 mm diameter filter paper punch,

and the extract was then moved away for further processing (i.e., derivatization and solvent

exchange, similar to Figure 3.4a). After wetting, the filter paper punch remains adhered to the

surface through capillary forces. Like method 1, this process requires ~50 min to complete.

Although analytes extracted from DBS samples (using conventional means) have been analyzed

by microfluidic methods,186 as far as we are aware, DMF method 2 is the first microfluidic

method that has been reported to accept a DBS punch directly as a sample input.

69

(a)

0

100

200

300

400

Extraction Solvent

3.2mm DBS

Punch DerivatizedExtract

1 2

(b)P

he

Co

nce

ntr

atio

n (µ

M)

Newborn Patient 1

Newborn Patient 2

Newborn Patient 3

120 µM

Figure 3.6: Analysis of AAs in DBS by DMF method 2. (a) Frames from a movie depicting sample processing of 3.2 mm diameter punch of a DBS on filter paper by DMF. (b) Graph of Phe concentrations measured by the DMF method in DBS punches from three NSO patients. As shown, patients 1 and 3 were correctly diagnosed as having phenylketonuria, and patient 2 was correctly identified as being unaffected. The dashed lines indicate the upper level for normal concentrations of Phe in newborn blood samples.

To evaluate DMF-driven method 2 relative to gold standard practices, a series of punches

from blood samples containing various concentrations of Phe were processed by DMF method 2,

and punches from the same samples were evaluated using the conventional newborn screening

technique. As listed in Table 3.3, a paired t-test revealed no significant difference between the

two data sets at a 95% confidence level. To validate the new technique for application to clinical

samples, DBS punches from three newborn patients of NSO were evaluated by the DMF method.

As shown in Figure 3.6b, the new technique correctly identified patients 1 and 3 as suffering

from phenylketonuria, and patient 2 as being unaffected.

70

Table 3.3. Measured phenylalanine (Phe) concentration in 3.2 mm dia. punches from filter paper bearing dried blood using DMF-driven method 2 (left) and standard techniques at NSO (right). The average difference between the measurements generated by the two methods is 6.4%.

Sample Measured Phe Concentration

(µM) Using DMF Method 2

Measured Phe Concentration (µM)

Using NSO Technique

1 70 70

2 550 548

3 93 88

4 93 92

5 368 302

6 534 539

7 735 871

3.3.3 DMF-Driven Method 3

While the data described above were generated with microfluidic sample processing and

off-line analysis, there may be some cases in which a fully integrated (processing + analysis)

method is useful. To accommodate this need, method 3 (Fig. 3.2c) was developed in which a

DMF platform was coupled directly to a nanoESI emitter for in-line analysis by MS. As shown

in Figure 3.3, the central feature of this method (building on recent work with hybrid

microfluidics108) is an intersection of a DMF electrode and a microchannel through a vertical

hole. In this geometry, droplets are manipulated on the top surface, and are subsequently

71

transferred to microchannels on the bottom of the device through the hole. A portion of an

experiment using method 3 is depicted in Figure 3.7a. A blood sample was first spotted on the

device as in method 1 (note that analogous methods could be developed using DBS sample

punches similar to method 2) and the AAs were extracted and derivatized as described above.

The dried, derivatized sample was then resuspended and the droplet was actuated to the access

hole such that it filled the channel below by capillary action. A nanoelectropsray is generated at a

corner of the device187,188 (Figure 3.7b) by applying a high voltage to the counter electrode.

Representative mass spectra generated from samples processed and analyzed on-chip are shown

in Figure 3.7c. The entire process requires ~1 h from sampling to analysis, and requires only the

hybrid DMF device and a mass spectrometer (i.e., no nanoflow pumps, capillary connections,

robots, or samplers).

72

Blood Sample

Extraction Solvent

DerivatizationSolvent

AccessHole

Empty Microchannel1

MS Solvent

DerivatizedExtract

2 Filled Microchannel

MS Solvent

3

MS Orifice

Spray

Chip Orifice

100 m/z

Inte

nsity

(arb

)

MS2 MS2125 120

150 200 250 100 150 200 250

m/z

(a)

(b) (c)

Figure 3.7: Analysis of AAs in DBS by DMF-driven method 3. (a) Frames from a movie (left-to-right) demonstrating derivatization and extraction of AA from dried blood, resolubilization in solvent, and analyte solution in spray microchannel. (b) Image of sample spraying from an integrated emitter. (c) MS2 spectra of Phe (left) and d5-Phe (right) generated from blood samples.

73

3.4 Discussion

Newborn screening programs have been implemented around the world to detect

aminoacidopathies at an early stage.166,167,189 MS/MS, which facilitates the simultaneous

quantification of multiple analytes, is particularly useful for such programs, but the technique is

hampered by disadvantages associated with sample preparation. Here, we report a set of three

new automated microfluidic methods powered by DMF for quantifying amino acids in blood

samples, and present proof-of-concept results demonstrating that these techniques may be useful

for screening samples for homocystinuria, phenylketonuria, and certain types of tyrosinemia. The

new methods have several advantages relative to the conventional techniques, which are

discussed below.

An obvious advantage of the microfluidic methods relative to conventional techniques is

automation -- if a robotic screening platform is not available, a single technician using an

instrument powered by DMF could likely do the work of several technicians using manual

techniques. Furthermore, the DMF-driven methods facilitate reduction in reagent use (20 µL vs.

170-450 µL164,190,191), and a reduction in analysis time (~1 h vs. >3.5 h170,171). Note that the 3.5 h

reported for the conventional method is for analysis of 96 samples, while the 1 h required by the

new method is for the analysis of 3 samples. We propose that future generations of the DMF

technique should be capable of accommodating 96 samples in parallel.

Another advantage of the new technique is the potential for direct integration with mass

spectrometry (Fig. 3.7). NanoESI is a finicky technique requiring operator expertise and

vigilance to achieve reproducible results, which is, in part, the reason why all newborn blood

samples in Ontario are mailed to a single facility.160 Indeed, a recent editorial on this subject

74

decried the inherent limitations of systems relying on “conventional electrospray plumbing

including capillary tubes etc”.170 The device shown in Figure 3.7 is the first ever reported to

integrate DMF with in-line MS. In this design, the nESI plumbing is built-in to the device by

standard batch-processing. Sample analysis is realized simply by positioning a device in front of

the mass spectrometer and applying an electrical potential.

Over the past two decades MS/MS has revolutionized newborn screening, replacing the

tests used historically for diagnosing amino acid disorders (i.e., Guthrie’s bacterial inhibition192

and fluorometric193 and enzymatic194 tests). This is mainly because MS/MS facilitates the

analysis of multiple analytes simultaneously in a single run in comparison to the traditional

single-analyte assays, which improves efficiency and reduces costs.163,170 In this work, we

present the first method to combine microfluidic sample handling and preparation with MS/MS

for analysis of amino acids. We note that there are other metabolic and non-metabolic disorders

(e.g., cystic fibrosis, congenital hypothyroidism, biotinidase deficiency, galactosemia etc.) that

are commonly screened in newborns that are not routinely detected by MS/MS. Instead, these

diseases are evaluated by enzyme-, DNA-, or immunoassay-based tests163,195 and a variety of

new technologies195,196 (including digital microfluidics197) are being developed to implement

them. We speculate that future microfluidic systems might be capable of implementing all of the

necessary tests, including those that rely on MS and those that do not.

The current work demonstrates proof-of-concept for quantification of three AAs (Met,

Phe, and Tyr) that are often measured for early diagnosis of diseases in newborn screening, and

in on-going work, we are extending the technique to be compatible with the full suite of diseases

tested by MS/MS by NSO.160 If successful, we propose that these methods might be useful for a

75

wide range of applications. For example, we propose that blood samples could be spotted

directly onto inexpensive devices86,87 (or removable device coverings106), after which they would

be returned to a laboratory for analysis. Alternatively, punches from DBS samples on filter paper

could be used (i.e., method 2), and either sampling technique might be combined with in-line

analysis (i.e., method 3). We posit that the new methods presented here represent an attractive

advance for the growing number of applications relying on DBS samples in the medical clinic

and the pharmaceutical industry.

76

3.5 Conclusion

We report new methods for sample processing and analysis of markers of amino acids in

dried blood samples using an integrated microfluidic device. The new methods are automated,

and facilitate significant reductions in reagent consumption. In addition, the new methods have

built-in plumbing for direct interfacing with mass spectrometry. We propose that these advances

have the potential to contribute to a new paradigm of fast, inexpensive screening.

77

Chapter 4 Digital Microfluidic Electrospray Ionization Interface for Succinylacetone Analysis in Dried Blood Spots

The multilayer hybrid microfluidic device described in Chapter 3 was the first method

developed to interface DMF to ESI-MS for inline sample analysis. That device requires an

access well to transfer the DMF-processed droplet from the top substrate to a bottom

microchannel emitter via capillary action. Although the results obtained using the hybrid devices

were promising, they required complex fabrication and bonding processes. Here we report a

simpler and more direct method for coupling DMF with ESI-MS. The new digital microfluidic

electrospray ionization (DMF-ESI) interface was formed by sandwiching a pulled glass capillary

emitter between the two DMF substrates (a bottom substrate bearing an array of actuation

electrodes and a top substrate bearing a counter electrode) with an inter-substrate gap selected

such that the capillary emitter is immobilized without any external seals or gaskets. As proof of

concept, on-chip processing and analysis of succinylacetone from dried blood spots was

performed. This chapter represents a snapshot of a work in progress; we anticipate that additional

work in the near future will complete the story.

78

4.1 Introduction Succinylacetone (SA) is a metabolite of tyrosine, and is a blood biomarker for

hepatorenal tyrosinemia (HT), a metabolic disorder characterized by life-threatening progressive

liver and kidney dysfunction and hepatocellular cancer.198 Hence, the ability to quantify this

analyte is critical, as treatment and management strategies can significantly reduce the morbidity

and mortality rate of this disease. In fact, SA measurements have replaced the older technique of

measuring tyrosine levels in newborn blood samples, as recent studies198,199 have demonstrated

that increased blood concentrations of tyrosine are neither specific nor sensitive enough to yield

HT diagnoses. The switch to using SA (instead of tyrosine) has significantly reduced the high

number of false positives and false negatives.162,199-202

Biomarkers for tyrosinemia and a host of other metabolic diseases are typically measured

from filter paper samples bearing dried blood spots (DBS).159 A variety of different methods

have been developed to quantification SA in such samples.203-206 For example, SA can be

extracted and derivatized by adding hydroxylamine hydrochloride204 or dansylhydrazine203 to a

DBS followed by LC-MS/MS analysis of the extractate. These methods are reproducible and

have sub-µM detection limits; however, these methods consume a native blood sample which

precludes the use of a single sample for multiple analyses. On the other hand, Allard et al.205 and

Sander et al.206 demonstrated an alternative technique in which SA is extracted and derivatized

from so-called residual blood spots that have been pre-extracted with absolute methanol. In this

method, the hydrazine reacts with SA to form a hydrazone derivative, which is selectively

soluble in the extraction solvent. This method is particularly advantageous because the use of

79

residual blood spots allows for a single sample to be used for multiple analyses (i.e., the pre-

extraction in methanol can be used to quantify amino acids, as described in chapter 3), and also

facilitates SA measurements with higher sensitivity (less matrix effects) as well as shorter

analysis time.

In the work described in Chapter 3, we demonstrated the utility of DMF-driven

processing of dried blood spot samples for quantification of amino acids. Here, as an extension

to the previous work, we describe a related method to extract and quantify SA from residual

blood spots on a DMF platform. The device represents an improvement over the method

described in Chapter 3 in that fabrication of the interface is much simpler -- a pre-formed pulled-

glass capillary emitter is mated directly to a two-plate DMF device (in a format similar to what

was described in a recent patent application207). In addition, the new method is fully automated,

using an impedance-based feedback mechanism for high-fidelity droplet actuation.208 This

chapter represents work in progress; however, the initial results are encouraging, and we propose

that the new device and method format described here may be useful for myriad applications in

which samples must be processed prior to analysis with mass spectrometry.

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4.2 Reagents and Materials 4.2.1 Study Subjects

Liquid blood samples were collected from a healthy adult male volunteer after a 10 h

fasting period, and were kept at –20°C until analysis.

4.2.2 Reagents and Materials

Unless otherwise specified, reagents were purchased from Sigma Chemical (Oakville,

ON). 3,4,5,6,7-13C-succinylacetone (513C-SA) was obtained from Cambridge Isotope

Laboratories (Andover, MA). WhatmanTM filter paper was purchased from Fisher Scientific

(Ottawa, ON). All working solutions were prepared using deionized (DI) water that had a

resistivity of 18 MΩ•cm at 25°C, filtered with nylon syringe filters from Millipore (Billerica,

MA, 0.2 µm pore diameter) and sonicated (5 min) prior to use.

For calibration, whole blood was fortified with SA (5, 10, 20, 50, 100 µM) and 513C-SA

(15 µM). 100 µL of each calibration sample was spotted on WhatmanTM filter paper and dried at

ambient temperature overnight. After drying, 2.5 mm diameter punches were generated using a

biopsy punch tool (Surgical Tools, Bedford, VA). These samples were stored at 4ºC prior to

analysis.

4.2.3 DMF Device Fabrication

Digital microfluidic devices were fabricated in the University of Toronto Emerging

Communications Technology Institute (ECTI) cleanroom facility, using a transparent photomask

printed at Pacific Arts and Design (Markham, ON). The bottom plates of DMF devices were

81

formed from glass substrates bearing patterned chromium electrodes by photolithography and

etching as described previously,176 and were coated with 7 µm of Parylene-C and 50 nm of

Teflon-AF. Parylene-C was applied using a vapor deposition instrument (Specialty Coating

Systems), and Teflon-AF was spin-coated (1% wt/wt in Fluorinert FC-40, 1000 rpm, 30 s)

followed by post-baking on a hot-plate (160ºC, 10 min). The polymer coatings were removed

from contact pads by gentle scraping with a scalpel to facilitate electrical contact for droplet

actuation. In addition to patterned bottom-plate substrates, unpatterned indium tin oxide (ITO)

coated glass substrates (Delta Technologies Ltd, Stillwater, MN) were coated with Teflon-AF

(50 nm, as above) for use a top-plate substrates.

Sample

DerivatizationReagent

Glass Capillary

Extraction Solvent

Processed sample

(a) (b)

(c) (d)

MS Orifice

nanospray

Gap360 µm

Top Plate

Bottom Plate

ElectrodeDielectric Glass

Hydrophobic

~~

Side view

Droplet Glass Tip

Top view

Figure 4.1: DMF-nanoESI interface. (a) Top-view schematic showing 7 reservoir electrodes for solvents and reagents, 22 DMF driving electrodes, and a pulled glass capillary inserted between two DMF substrates. (b) Side-view schematic. (c) Image of a device showing an array of contact pads on the side used to mate with a 40-pin connector for automated droplet control. (d) Image of spray generated off the tip of the capillary.

82

The device shown in Figure 1 was used for automated DMF-driven DBS processing and

analysis. It featured an array of twenty-two actuation electrodes (2.2 x 2.2 mm ea.) connected to

7 reservoir electrodes (5 x 5 mm ea.), with inter-electrode gaps of 40 µm. Each driving electrode

and reservoir was connected to a contact pad in an array of pads on the side of the device spaced

appropriately to interface with 40-pin connector (Compar Inc., Burlington ON). Devices were

assembled with an unpatterned ITO–glass top plate and a patterned bottom plate separated by a

spacer formed from four pieces of double-sided tape (total spacer thickness ~360 µm). With

these dimensions, the 10 µL droplets used for most experiments covered approximately five 2.2

x 2.2 mm electrodes.

4.2.4 DMF Device Operation

DMF droplet motion was managed using an automated feedback control system. In this

method, droplet movement is monitored by impedance sensing such that if a movement failure is

observed, additional voltage pulses are applied until the droplet completes the desired operation.

The system used here represents a new generation of an older system that we described (in

detail) previously.208 The differences between the two systems are summarized in Table 4.1.

Highlights of the new system include a new active impedance measurement circuit (shown in

Figure 4.2) and an improved feedback mechanism relying on progressive increases in driving

voltages (described in Table 4.1).

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Table 4.1: Differences between the new feedback control system used here and the feedback control system described previously.208

Property System Used Here System Used Previously208

Feedback voltage measurement circuit

Buffered system, shown in Figure 4.2, managed by an RBBB Arduino microcontroller (Modern Device, Providence, RI)

Passive system relying on resistors and capacitors

Circuit tuning mechanism

Digital potentiometer (Radj = 1- 5 k) managed by Arduino capable of automatic thresholding for new liquids during a given experiment

Manual potentiometer (Radj = 1- 20 k) which must be adjusted for each new liquid prior to each experiment

High voltage source and degree of control

Sine-wave from function generator (managed by software) and amplifier which can be adjusted during each experiment

Sine-wave from stand-alone function generator and amplifier which must be adjusted prior to each experiment

Length of each droplet actuation pulse

505 ms (500 ms driving pulse + 5 ms measurement)

215 ms (200 ms driving pulse + 15 ms measurement)

Control software and interface

Custom C++ program interfaced to Arduino microcontroller via USB cable

Custom LabView (National Instruments, Austin, TX) program interfaced to a DAQPAD 6507 (National Instruments)

Number, type, and configuration of relays/switches

80 AQV259H (Panasonic) solid-state relays - 2 per output (total 40 outputs), with each output capable of two states: high-voltage, ground

96 RT424012F (Tyco Electronics) mechanical relays - 2 per output (total 48 outputs) with each output capable of three states: high-voltage, ground, and float

Feedback mechanism

Repeated pulses with a 50 VRMS step-up in voltage every second (up to a maximum of 350 VRMS) until droplet movement is achieved

Repeated pulses of the same actuation voltage until droplet movement is achieved

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4.2.5 DMF-ESI-MS Interface

After assembling the DMF device, a ~5 cm long, 360 μm O.D., 50 μm I.D. pulled glass

capillary ESI emitter (New Objective Inc., Woburn, MA) was carefully inserted between the two

plates of the DMF device. The device was then positioned such that the tapered tip of capillary

was ~3 mm away from the orifice of MS (Figure 4.1d). To initiate analysis by mass

spectrometry, a droplet was driven to the edge of the pulled-glass emitter, and after filling by

capillary action, 1.7-2.2 kV was applied to the ITO-coated top plate of the DMF device to

generate a nanoelectrospray into an LTQ Mass Spectrometer (Thermo Scientific). A solution of

tyrosine (5 µM in 50:50 MeOH:H2O) was used to evaluate the efficacy of the system for ESI-MS

analysis.

4.2.6 DMF-driven SA Processing

Prior to DMF actuation, a 2.5 mm dia. DBS punch was positioned on the bottom plate of

a device. The top plate was then affixed and extraction solvent (MeOH) was loaded in the

appropriate reservoir. Five sequential steps were then implemented. First, a 10 µL droplet of

MeOH was dispensed from the appropriate reservoir and actuated toward the DBS punch. After

15 minutes of incubation at room temperature, the extraction solvent was actuated away from the

DBS and collected in the waste reservoir. Next, a 10 µL droplet of derivatization solvent (20 µM

N2H4 in 80:20 ACN:H2O) was dispensed to the residual blood spot. After incubating the reaction

for 15 min at 37ºC, a fresh 10 µL droplet of derivatization solvent was dispensed and delivered

to the residual blood spot, after which it was actuated to the capillary emitter positioned at the

edge of the electrodes.

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4.2.7 SA Quantification by NanoESI-MS/MS

Tandem MS/MS analysis of SA extracted from blood spots was carried out by

introducing 25% collision energy to the parent ions and then the fragments over the m/z range of

100-300 were scanned. SA daughter ions, which exhibit a loss of H2O (18 m/z), were observed in

the second mass selection and the peak intensities were used for quantification. Spectra were

collected as an average of 50 acquisitions, and at least three samples were evaluated for every

condition recorded. Calibration curves were generated by plotting the intensity ratios of the

daughter ions of SA relative to internal standard (5C13 SA) as a function of SA concentration.

86

4.3 Results and Discussions

4.3.1 DMF Fabrication and Operation

As shown in Figure 1, the automated DMF design included twenty-two driving electrodes

and seven reservoir electrodes. The device features an array of contact pads patterned at the edge

of the device, which served as the interface between the device and the automated feedback

control system (Figure 4.2). For many liquids, droplet movement was observed to be facile and

feedback control was not important (i.e., in many cases, every voltage pulse resulted in

successful droplet movement). But for the case of manipulating droplets after they had touched

punched filter paper, droplet movement was observed to be sluggish and inconsistent. In this

case, feedback control was critical to ensure successful completion of the programmed sequence.

87

Figure 4.2: Schematic of the feedback control system, showing the relationships between the function generator and amplifier, control board, RBBB Arduino microcontroller, the DMF device, the PC, and the feedback circuit. To actuate an electrode, the user clicks an electrode in the C++ interface, which sends a 5 Vpp signal to the Arduino to activate the designated high voltage relay on the control board. The feedback circuit detects Vfeed and compares it with a threshold to determine whether the droplet movement was successful. A list of differences between this system and one described (in detail) previously208 is found in Table 1.

4.3.2 DMF-ESI-MS Interface

As shown in Figure 4.1 the new DMF-ESI-MS interface was constructed by inserting a

pulled-glass capillary in between the two substrates of an assembled DMF device. The gap

between the two substrates was formed from 4 layers of double sided tape such that it was

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roughly equivalent to the outer diameter of glass capillary. With this configuration, capillary

emitters were easily introduced and replaced without requiring that the device be disassembled.

In contrast to the method described in Chapter 3 (with DMF electrodes mated to microchannels

through a vertical interface), in the new technique, the interface is decoupled from the DMF

device, which allows for multiple emitters to be used with no worry about cross contamination

arising from sample sticking to an emitter. In addition, this decoupling eliminated extra

fabrication steps and tedious alignment steps prior to thermal bonding of the two substrates.

Finally, the new system has improved overall efficiency in which failure in one part of the

platform (i.e., clogging of the tip or surface fouling on the DMF device) does not compromise

the efficiency of the entire system.

Once assembled, the device with connectors attached was mounted close to the orifice of

a mass spectrometer. After spontaneous filling of the emitter by capillary action, a spray was

generated via application of high voltage to the top-plate ITO electrode. A representative mass

spectrum of a solution of tyrosine is shown in Figure 4.3a. The consumption rate was estimated

to be ~800 nL/min for a 2 µL droplet, and the spray was stable for ~150 seconds with 8.6%

relative standard deviation (Figure 4.3b).

89

Time (s)

Intensity (arb)

Spray of 2 µL Tyrosine droplet

m/z

Intensity (arb)

200180160140120100

8.6 % RSD

5 µM Tyrosine

[M+H]+

(a)

(b)

140120100806040200

Figure 4.3: Performance of the new DMF-nanoESI interface. (a) MS1 spectrum of tyrosine. (b) Spray stability over the course of 150 seconds.

4.3.3 DMF-Driven SA Processing and Quantification

An automated digital microfluidic method was developed to analyze SA from DBS

samples on filter paper. A portion of an experiment is shown in Figure 4.4. First, a droplet of

MeOH was dispensed and driven to a 2.5 mm diameter punch from filter paper bearing a DBS

90

(Frame 1). After incubating for 15 min at room temperature (Frame 2), the extract was moved

away (Frame 3). The residual blood spot was further extracted and derivatized in a droplet of

N2H4 in 80:20 ACN:H2O (Frame 4), which was allowed to incubate for 15 minutes. After

extraction and derivatization, the extract droplet was actuated to the tip for spray into the mass

spectrometer (Frame 5). The entire process is fully automated and requires ~40 min to complete.

MeOH

1 3

5

Incubate

2Capillary

H4N2

4

DBS punch

Figure 4.4: Sequence of frames from a movie depicting SA extraction from a punch from a dried blood spot (DBS) by DMF. (1) Dispense a droplet of MeOH and actuate toward the DBS punch. (2) Incubate the MeOH droplet on the DBS punch. (3) Actuate away the MeOH droplet away from the DBS punch for collection in the designated reservoir. (4) Dispense a droplet of N2H4 extraction solvent to the residual blood spot. (5) Actuate the processed droplet toward the capillary tip.

The new method reported here comprises two different extractions -- into methanol

(frames 1-3 in Figure 4.4) and into a solution of hydrazine (frames 4-5 in Figure 4.4). This two-

step extraction procedure confers multiple advantages. First, although not implemented here, the

methanol extract droplet could be analyzed to quantify amino acids, acylcarnitines, and other

small, polar molecules (as in the work described in Chapter 3). Second, the pre-extraction in

methanol serves as a sample clean-up step, which is critical to the second extraction in

91

hydrazine. In preliminary work without an initial methanol extraction, the hydrazine reaction was

observed to produce a large number of (presumably non-SA-related) derivatives with poor

solubility that interfered with the analysis and made droplet movement sluggish and

unpredictable (even when using the feedback control system).

M/Z

2001801601401201008060

Intensity (arb)

MS2 SA (m/z 155 → 137)

2001801601401201008060

MS2 5C13 SA (m/z 160 → 142)

M/Z

Intensity (arb)

(a)

(b)

[M+H]+

[M+H‐H2O]+

[M+H‐H2O]+

[M+H]+

Figure 4.5: Tandem MS spectra of (a) derivatized SA and (b) 5C13 SA showing the loss of 18 as result of CID.

92

Typical MS/MS data from derivatized SA and its internal standard measured using the

DMF method in a residual blood spot is shown in Figure 4.5. When the product ions were

subjected to CID, a characteristic fragmentation (neutral loss of 18) was observed (i.e., m/z

155→137 for SA and 160 → 142 for 513C-SA). These transitions were used to quantify SA, and

a calibration curve was generated with R2 greater than 0.998 (Figure 4.6). In the near future, we

plan to use this method to quantify SA concentration in DBS punches bearing patient samples

from our collaborators at the Children’s Hospital of Eastern Ontario.

120100806040200

Rat

io o

f Int

ensi

ties

(SA

/5C

13SA

)

Succinylacetone (µM) Figure 4.6: Calibration curve generated by the DMF method for quantification of spiked SA in dried blood samples. Data were generated by plotting the intensity ratios of the daughter ions of SA relative to internal standard (5C13 SA) as a function of SA concentration. Each data point represents at least four replicate measurements, and error bars represent ±1 S.D. The calibration curve had an R2 of 0.998.

93

4.4 Conclusion

We have developed an automated microfluidic device with on-chip coupling of with a

nanoelectrospray emitter that facilitates inline analysis of DMF processed samples by mass

spectrometry. The new method incorporates a feedback control system enabling facile, high-

fidelity droplet movement without any manual intervention. As proof of principle, the method

was used to perform on-chip extraction and quantification of succinylacetone, a marker of

tyrosinemia, in dried blood spot samples. We propose that the new system represents a potential

prototype for fast and inexpensive screening of biomarkers for disease in dried blood spots and

other complex samples.

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Chapter 5 A World­to­Chip Interface for Digital Microfuidics DMF is a fluid handling technique that enables manipulation of discrete droplets on an

array of electrodes. There is considerable enthusiasm for this method because of the potential for

array-based screening applications. A limitation for DMF is non-specific adsorption of reagents

to device surfaces. If a given device is used to actuate multiple reagents, this phenomenon can

cause undesirable cross-contamination. A second limitation for DMF (and all other microfluidic

systems) is the “world-to-chip” interface; it is notoriously difficult to deliver reagents and

samples to such systems without compromising the oft-hyped advantages of rapid analyses and

reduced reagent consumption. In response to these limitations, we introduce a new strategy for

DMF, in which a removable plastic “skin” is used to (a) eliminate cross-contamination, (b)

bridge the world-to-chip interface, and (c) interface with MALDI-MS. We demonstrated the

utility of this format by implementing on-chip protein digestion on immobilized enzyme depots.

This new method has the potential to transform DMF from being a curiosity for aficionados into

a technology that is useful for biochemical applications at large.

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5.1 Introduction

There is currently great enthusiasm for applying DMF to a wide variety of applications

including cell-based assays,97,98 enzyme assays,88-90 protein sample preparation,91-96 and the

polymerase chain reaction.114 Unfortunately, there are two critical limitations on the scope of

applications compatible with DMF – biofouling and interfacing. The former limitation,

biofouling, is a pernicious one in all microscale analyses; a negative side-effect of high surface

area to volume ratios is the increased rate of adsorption of analytes from solution onto solid

surfaces. Luk105 and others88,209 have developed strategies to limit the extent of biofouling in

digital microfluidics, but the problem persists as a roadblock, preventing wide adoption of the

technique.

The second limitation for DMF (and for all microfluidic systems) is the “world-to-chip”

interface; it is notoriously difficult to deliver reagents and samples to such systems without

compromising the oft-hyped advantages of rapid analyses and reduced reagent

consumption.168,210 A solution to this problem for microchannel-based methods is the use of pre-

loaded reagents. Such methods typically comprise two steps: (1) reagents are stored in

microchannels (or in replaceable cartridges), and (2) at a later time, the reagents are rapidly

accessed to carry out the desired assay/experiment. Two strategies have emerged for

microchannel systems; in the first, reagents are stored as solutions in droplets isolated from each

other by plugs of air211 or an immiscible fluid59,212 until use. In a second, reagents are stored in

solid phase in channels, and are then reconstituted in solution when the assay is performed.213-215

Pre-loaded reagents in microfluidic devices is a strategy that will be useful for a wide range of

96

applications.216 Until now, however, there has been no analogous technique for digital

microfluidics.

In response to the twin challenges of non-specific adsorption and world-to-chip

interfacing in digital microfluidics, we have developed a new strategy relying on removable

polymer coverings.87,217 We call these coverings “skins” to highlight the central concept – after

each experiment, a thin film is replaced, but the central infrastructure of the device is reused. The

skins effectively prevent cross-contamination between repeated analyses, and perhaps more

importantly, serve as a useful medium for reagent introduction onto DMF devices. To

demonstrate this principle, we pre-loaded dried spots of enzymes to skins (which we call “skin

depots”) for subsequent use in proteolytic digestion assays. The loaded reagents were found to be

active after >1 month of storage in a freezer. As the first technology of its kind, we propose that

this innovation may represent an important step forward for digital microfluidics, making it an

attractive fluid-handling platform for a wide range of applications.

97

5.2 Materials and Methods

5.2.1 Reagents and Materials

Angiotensin I and II, bradykinin, bovine alpha-chymotrypsin and trypsin, proteinase K,

myoglobin, ubiquitin, rhodamine B, Tris-HCl, CaCl2, trifluoroacetic acid (TFA), Pluronic F127,

and acetonitrile were purchased from Sigma-Aldrich Canada (Oakville, ON). 20mer DNA

oligonucleotide (5′AGCAGAGCGACCT-CAATGAT3′) was purchased from Sigma Genosys

(Oakville, ON). An Enzchek protease assay kit (including labeled, quenched BODIPY-casein)

and FITC-labeled bovine serum albumin (FITC-BSA) were obtained from Invitrogen

(Burlington, ON). Ultramarker was obtained from Thermo Fisher Scientific (Ottawa, ON).

Alpha-cyano-4-hydroxycinnamic acid (α-CHCA), 2,5-dihydroxybenzoic acid (DHB), 3-

hydroxypicolinic acid (HPA) and sinapinic acid (SA) were purchased from Waters Limited

(Mississauga, ON). Parylene-C dimer was from Specialty Coating Systems (Indianapolis, IN),

and Teflon-AF was from DuPont (Wilmington, DE). 1-Ct silicone oil (vapor pressure = 4 mm

Hg at 25ºC) was purchased from Gelest Inc. (Morrisville, PA). Polyethylene food wrap with

trademark “Saran” was purchased from grocery stores, and plastic tape was from Grand & Toy

(Toronto, ON).

Clean room reagents and supplies included Shipley S1811 photoresist and MF321

developer from Rohm and Haas (Marlborough, MA), AZ300T photoresist stripper from AZ

Electronic Materials (Somerville, NJ), solid chromium from Kurt J. Lesker Canada (Toronto,

ON), CR-4 chromium etchant from Cyantek (Fremont, CA), and conc. sulfuric acid and

98

hydrogen peroxide (30%) from Fisher Scientific Canada (Ottawa, ON). Piranha solution was

prepared as a 3:1 (v/v) mixture of sulfuric acid and hydrogen peroxide.

Working solutions (10 mg/mL) of all matrixes (α-CHCA, DHB, HPA, and SA) were

prepared in 50% analytical grade acetonitrile/deionized (DI) water (v/v) and 0.1% TFA (v/v) and

were stored at 4°C away from light. Stock solutions (10 µM) of angiotensin I, II and bradykinin

were prepared in DI water, and stock solutions (100 µM) of ubiquitin and myoglobin were

prepared in working buffer (10 mM Tris-HCl, 1 mM CaCl2 0.0005% w/v Pluronic F127, pH 8).

All stock solutions of standards were stored at 4°C. Stock solutions (100 µM) of digestive

enzymes (bovine trypsin, α-chymotrypsin, and proteinase K) were prepared in working buffer

and were stored as aliquots at -80°C until use. Immediately preceding assays, standards and

enzymes were warmed to room temperature and diluted in DI water (peptides) or working buffer

(proteins, enzymes, and fluorescent reagents). Fluorescent assay solution (3.3 µM quenched

BODIPY-casein and 2 µM rhodamine B in working buffer) was prepared immediately prior to

use.

5.2.2 Device Fabrication and Operation

Devices were fabricated in the University of Toronto Emerging Communications

Technology Institute (ECTI) fabrication facility using processes similar to those reported

previously.87,105 Briefly, glass slides were cleaned with piranha solution (15 min), and then

coated with 200 nm of chromium by electron beam deposition. After rinsing and baking on a hot

plate (110°C, 5 min), the substrates were primed by spin-coating with HMDS (3000 RPM, 30 s),

and were then spin-coated with Shipley S1811 photoresist (3000 RPM, 30 s). Substrates were

pre-baked on a hotplate (100°C, 5 min), and exposed through a photomask using a Suss

99

Mikrotek mask aligner. Substrates were developed in MF321 (3 min), and then post-baked on a

hot plate (100°C, 3 min). After photolithography, substrates were immersed in chromium etchant

(1 min). Finally, the remaining photoresist was stripped in AZ300T (10 min).

Prior to experiments, devices were fitted with un-modified polymer coatings formed from

food wrap (called “skins”), or reagent-loaded polymer coatings (called “skin-depots”). Un-

modified skins were prepared by adhering coverings to unpatterned glass substrates, spin-coating

with Teflon-AF, (1% w/w in Fluorinert FC-40, 1000 RPM, 60s), and post-baking on a hot plate

(75°C, 30 min). When ready for use on digital microfluidic devices, a few drops of silicone oil

were dispensed onto an electrode array, followed by application of the skin (Teflon-AF side up),

followed by annealing on a hot plate (75°C, 2 min). Skin-depots were prepared as above, but

prior to use, they were modified by application of reagent spots. This step was achieved by

pipetting 2 µL droplet(s) of enzyme (6.5 µM trypsin,10 µM α-chymotrypsin, or 13 µM proteinase

K) onto the Teflon-coated surface, and allowing them to dry. Skin depots were either used

immediately, or were sealed in a plastic Petri-dish and stored at -20°C or -80°C. Prior to use,

skin depots were allowed to warm to room temperature (if necessary), peeled off of the

unpatterned substrate, applied to a silicone-oil coated electrode array, and annealed on a hot plate

(75°C, 2 min). In addition to food wrap, plastic adhesive tape and wax film (Parafilm M, Alcan

Packaging, Neenah, WI) were also evaluated for use as digital microfluidic device skins. Skins

formed from tape were attached to devices by applying gentle pressure, while those formed from

wax film were manually stretched (to about 10 µm thickness) and then wrapped around the

device to make a tight seal free of air bubbles.

100

Devices had a “Y” shape design of 1 mm × 1 mm electrodes with inter-electrode gaps of

10 µm. 2 µL droplets were moved and merged on devices operating in open-plate mode (i.e.,

with no top cover) by applying driving potentials (400-500 VRMS) to sequential pairs of

electrodes. The driving potentials were generated by amplifying the output of a function

generator operating at 18 kHz, and were applied manually to exposed contact pads. Droplet

actuation was monitored and recorded by a CCD camera.

5.2.3 Mass Spectrometry

Matrix assisted laser desorption/ionization mass spectrometry (MALDI-MS) was used to

evaluate samples actuated on DMF devices. Matrix/sample spots were prepared in two modes:

conventional and in situ. In conventional mode, samples were manipulated on a device, collected

with a pipette and dispensed onto a stainless steel target. A matrix solution was added, and the

combined droplet was allowed to dry. In in situ mode, separate droplets containing sample and

matrix were moved, merged, and actively mixed by DMF, and then allowed to dry onto the

surface. In in situ experiments involving skin depots, matrix/crystallization was preceded by an

on-chip reaction: droplets containing sample proteins were driven to dried spots containing

digestive enzyme. After incubation with the enzyme (room temperature, 15 min), a droplet of

matrix was driven to the spot to quench the reaction and the combined droplet was allowed to

dry. After co-crystallization, skins were carefully peeled off of the device, and then affixed onto

a stainless steel target using double-sided tape. Different matrixes were used for different

analytes: α-CHCA for peptide standards and digests, DHB for ultramarker, HPA for

oligonucleotides and SA for proteins. At least three replicate spots were evaluated for each

sample.

101

Samples were analyzed using a MALDI-TOF Micro-MX MS (Waters, Milford, MA)

operating in positive mode. Peptide standards and digests were evaluated in reflectron mode over

a mass to charge ratio (m/z) range from 500-2,000. Proteins were evaluated in linear mode over a

m/z range from 5,000-30,000. At least one hundred shots were collected per spectrum, with laser

power tuned to optimize the signal to noise ratio (S/N). Data were processed by normalization to

the largest analyte peak, baseline subtraction, and smoothed with a 10-point running average.

Spectra of enzyme digests were analyzed using the Mascot protein identification package

searching the SwissProt database. The database was searched with 1 allowed missed cleavage, a

mass accuracy of +/- 1.2 Da, and no further modifications.

5.2.4 Fluorescence

Confocal fluorescence microscopy was used to evaluate protein adsorption on surfaces,

using methods similar to what Luk reported previously.105 Briefly, before and after a droplet

containing FITC-BSA (7 µg/ml) was driven across a device (4 times), the surface of the skin was

imaged using a Fluoview 300 scanning confocal microscope (Olympus, Markam, ON) equipped

with an Ar+ (488 nm) laser, in conjunction with a 10× objective (N.A. 0.95) (Fig. 5.1a).

Fluorescence was passed through a 510-525 nm band-pass filter, and each digital image was

formed from the average of four frames using FluoView image acquisition software (Olympus).

A fluorescence plate reader was used for long-term skin depot stability assays. Skin

depots were thawed and affixed onto devices, and a 2 µL droplet containing the assay reagent

(quenched BODIPY-casein) and internal standard (IS) (rhodamine B) was driven to the dried

spot by DMF. After reaction for 30 minutes in a humidified chamber, the device was positioned

on the top of a 96-well plate (with the droplet aligned to one of the wells) and inserted into a

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PheraStar multiwell plate reader (BMG Labtech, Durham, NC). The droplet was evaluated by

sequential measurements for BODIPY-casein signal (λex = 485 nm; λem = 520 nm) and IS signal

(λex = 530 nm; λem = 600 nm). Five replicates (using five separate skin depots) were evaluated for

each storage period (1, 2, 3, 10, 20, or 30 days), and the data were evaluated for statistical

significance using a two-tailed student’s t-test.

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5.3 Results and Discussion

5.3.1 Protein Adsorption and Cross Contamination

As is the case for most substrates (e.g., pipette tips, centrifuge tubes, multiwell plates,

etc.), digital microfluidic devices are prone to non-specific adsorption of proteins. For example,

as shown in Figure 5.1a, after a droplet containing FITC-BSA is translated on a DMF device, a

residue is left on the surface that can be detected by confocal microscopy. Such residues can

cause two types of problems for DMF: (1) the surface may become sticky, which impedes

droplet movement, and (2) if multiple experiments are to be performed, cross-contamination may

be a problem.

An example of evidence for cross-contamination on a DMF device is shown in

Figure 5.1(b-c). In this experiment, two different analytes (10 µM angiotensin I in the first run, 1

µM angiotensin II in the second) were evaluated by MALDI-MS after actuation across the same

path on the same device. As shown, the spectrum of angiotensin I generated after the first run

(Fig. 5.1b) is clean; however, the spectrum of angiotensin II (Fig. 5.1c) is contaminated with

residue from the previous run. In these tests, after actuation by DMF, the sample droplets were

transferred to a MALDI target for crystallization and analysis, meaning that the cross-

contamination comprised both (a) an adsorption step from the droplet to the surface in the first

run, and (b) a desorption step from the surface to the droplet in the second run. These processes

must occur rapidly, as the droplets are in contact with the surface for a short period of time. This

unpredictable and rapid adsorption and desorption (and the ensuing cross-contamination) was the

initial motivation for the work reported here.

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BEFORE MovementBEFORE Movement

AFTER MovementConfocal Microscopy

Images

FITC-BSA

1mm

Angiotensin II

cross-contamination from Angiotensin I

Angiotensin I

(b)

(c)

(a)

1st run

2nd run

Figure 5.1: Images and spectra demonstrating protein adsorption and cross-contamination in digital microfluidics. In (a), the top image shows a device prior to droplet actuation, paired with a corresponding confocal fluorescence image of a central electrode. The bottom image shows the same device after a droplet containing FITC-BSA (7 µg/mL) has been cycled over the electrode 4 times, paired with a corresponding confocal image. The two fluorescence images were processed identically. The spectra in (b) and (c) were generated from two different droplets manipulated on the same digital microfluidic device (the first and second runs, respectively). The spectrum generated in the first experiment (b) is of 10 µM angiotensin I (MW 1296); the spectrum generated in the second experiment (c) is of 1 µM angiotensin II (MW 1046). In (c), cross-contamination from the first run is observed.

5.3.2 Digital Microfluidic Skins

To overcome the phenomenon of non-specific adsorption, we developed a new strategy

for DMF, which facilitates unlimited use of a given device. As depicted in the cartoon in Figure

5.2, this strategy comprises replacing the permanent insulating layer used in conventional DMF

devices with a removable “skin.” In this format, reactions and assays are carried out as usual, by

applying sequences of electrical potentials to merge and mix reagent droplets. As is the case for

conventional DMF, reagent molecules tend to adsorb to the surface (depicted in Figure 5.2, step

3, as colored dots). After the assay is complete, the skin can be peeled off from the device and

105

replaced with a fresh one for subsequent use. Thus, the device can be used indefinitely by

eliminating cross-contamination. In addition, for assays that involve the formation of dried

products (e.g., co-crystallization with matrix for MALDI-MS93-95), the skins serve as a

convenient analysis tool – the product of a given assay can be analyzed while at the same time,

the device is re-used for subsequent assays.

device

reagent Askin

reagent B

residue

residueproduct

peel off

attach

analyze product

(1)

(3)

(4)

(2)

Figure 5.2: Schematic depicting the removable skin strategy. (1) Fresh piece of polymer covering is affixed to a DMF device; (2) reagent droplets are actuated on top of the covering to facilitate mixing and merging; (3) residue is left behind as a consequence of non-specific adsorption of analytes; (4) the plastic covering is peeled off and the product can be analyzed if desired (a picture of a device with a skin formed from adhesive tape is shown in the bottom right). This process can be repeated ad nauseum by using additional skins.

To illustrate the new strategy, four different types of analytes were processed on a single

DMF device, using a fresh skin for each run. As shown in Figure 5.3, the four analytes included

insulin (MW 5733), bradykinin (MW 1060), a 20-mer oligonucleotide (MW 6135), and the

synthetic polymer, Ultramark 1621 (MW 900-2200). Each skin was analyzed by MALDI-MS in-

situ, and no evidence for cross-contamination was observed. In the work described here, a

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relatively simple Y-shaped electrode design was used (suitable for one-step processes); however,

we also confirmed that the skin strategy is compatible with more complex electrode arrays

appropriate for multistep reactions. This is important, because the more complex the design, the

more care is required in fabrication. In my lab, DMF devices are typically disposable (used once

and then discarded); however, in experiments with removable skins, we regularly used devices

for 9-10 assays with no drop-off in performance. Thus, in addition to eliminating cross-

contamination, the skin strategy significantly reduces the fabrication load required to support

DMF.

[M + Na]+

[M + H]+

[M + Na]+

[M + 2Na]2+

(a)

(c) (d)

(b)

[M + H]+

Insulin Bradykinin

Oligonucleotide Ultramark

Figure 5.3: MALDI-MS analysis of different analytes processed on different skins using a single DMF device. (a) 35 µM insulin (MW 5733); (b) 10 µM bradykinin (MW 1060); (c) 10 µM oligonucleotide (MW 6135); (d) 0.01% ultramark (MW 900-2200).

107

In these experiments, we evaluated three different types of substrates for forming skins –

polyethylene food-wrap sheets (~15 µm thick), clerical adhesive tape (~45 µm) and stretched

sheets of wax film (~10 µm). Each of these materials was found to support droplet movement

and facilitate device re-use. Skins formed from food-wrap were the most reliable, and were used

to collect the data reported here; however, a drawback for this material is the requirement of

thermal annealing prior to use. In contrast, skins formed from tape and wax films did not require

an annealing step, which made them convenient to use. Other concerns, however, made these

materials less attractive. Skins formed from adhesive tape tend to damage the actuation

electrodes after repeated applications (although this would likely not be a problem for low-tack

tapes, which were not tested here). In addition, the thick tape substrates required the application

of larger driving potentials (~900 VRMS) for droplet manipulation. In contrast, the thin stretched

wax films were compatible with driving potentials similar to those used for skins formed from

food wrap. However, the thickness of skins formed in this manner was observed to be non-

uniform, making the skins less reliable for droplet movement. Regardless, these preliminary

results suggest that a variety of different materials (e.g., thin films of silicone or other plastics)

may be compatible with the removable skin concept.

Two potential pitfalls for the removable skin strategy are trapped air bubbles and material

incompatibility. In initial experiments with skins formed from food-wrap, bubbles were

occasionally observed to become trapped between the skin and the device surface during

application. When a driving potential was applied to an electrode near a trapped bubble, arcing

was observed, which damaged the device. We found that this problem could be overcome by

moistening the device surface with a few drops of silicone oil prior to application of the plastic

108

film. Upon annealing, the oil evaporates, leaving a bubble-free seal. The latter problem, material

incompatibility, might be a concern if materials in the skin were found to leach into solution,

which could interfere with assays. The Teflon coating makes this unlikely, and in my

experiments, no contaminant peaks were observed in any MALDI-MS spectra (including in

control spectra generated from bare skin surfaces, not shown). We cannot rule out the possibility

of this being a problem in other settings, but given the apparent wide range of materials that can

be used to form skins (see above), we are confident that alternatives could be used in cases in

which Teflon-coated polyethylene food wrap is not tenable.

5.3.3 Skin Depots

In exploring the replaceable skin strategy to overcome fouling and cross-contamination,

we realized that this method could, in addition, serve as the basis for an exciting new innovation

for digital microfluidics. By pre-depositing reagents onto skins (and by having several such

substrates available), the skins are transformed into a convenient new platform for rapid

introduction of reagents to a device. Thus, this “skin depot” strategy is a potential solution to the

well-known world-to-chip interface problem for microfluidics.168,210

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Substrate

MALDI matrixenzyme

depot

-chymotrypsin depot

Trypsin depot*

*

* * *

* * *

*

*

* = Predicted Peptide

Figure 5.4: Schematic and spectra demonstrating the skin depot strategy. The spectra were generated from ubiquitin samples digested on pre-spotted trypsin (top) and α-chymotrypsin (bottom). Peaks labelled with asterisks (*) were identified via a database search using Mascot, and the sequence coverage for all analyses was found to be greater than 50%.

110

(a)

Trypsindepot

-chymotrypsindepot

Proteinase K depot

(b)trypsin -chymotrypsin

proteinase K

myoglobin1.

2.

3.matrix crystal

(a)

Trypsindepot

Trypsindepot

-chymotrypsindepot

-chymotrypsindepot

Proteinase K depot

Proteinase K depot

(b)trypsin -chymotrypsin

proteinase K

myoglobin1.

2.

3.matrix crystal

Figure 5.5: Multiplexed skin depot. (a) A series of pictures from a movie (top-to-bottom) demonstrating a skin depot analysis in parallel. Droplets of myoglobin (panel 1) were delivered to enzyme depots and allowed to incubate (panel 2), followed by delivery of matrix and drying onto the skin (panel 3). (b) Spectra were generated from myoglobin samples digested on pre-spotted trypsin (top), α-chymotrypsin (middle) and proteinase K (bottom). Peaks labelled with asterisks (*) were identified via a database search using Mascot, and the sequence coverage for all analyses was found to be greater than 50%.

To illustrate the new strategy, we prepared skins loaded with dry enzyme depots, and

then used DMF to deliver droplets containing substrates and MALDI matrixes to the spots for

analysis. In one type of experiment, illustrated in Figure 5.4, each skin was loaded with a single

enzyme, and multiple assays were carried out sequentially by exchanging the skins. This strategy

would be appropriate for cases in which an end-user wants to rapidly switch between different

analyses for different samples. In a second type of experiment, illustrated in Figure 5.5, each skin

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was loaded with multiple enzymes, and assays were carried out in parallel. This strategy would

be appropriate for cases in which an end-user wants to carry out multiplexed assays on a given

sample. While these proof-of-principle experiments comprise a small number of reagents, we

speculate that in the future, a microarray spotter could be used to form skins appropriate for

high-throughput analysis. Regardless, the performance of all of the proteolysis assays reported

here was excellent; when evaluated using the proteomic search engine Mascot, sequence

identification was 50% or more for all cases. In optimizing the skin depot strategy for protease

assays, we observed the method to be quite robust despite a number of potential pitfalls. First,

pluronic F127 was used as a solution additive to facilitate movement of the analyte droplets (in

this case, ubiquitin and myoglobin); this reagent has been shown to reduce ionization efficiencies

for MALDI-MS.218 Fortunately, the amount used here (0.0005% w/v) was low enough such that

this effect was not observed. Second, enzymatic autolysis peaks were only rarely observed,

which we attribute to the low enzyme-to-substrate ratio and the short reaction time. Third, in

preliminary tests, we determined that the annealing step (75°C, 2 min) did not affect the activity

of dried enzymes. In the future, if reagents sensitive to these conditions are used, we plan to

evaluate skins formed from materials that do not require annealing (such as low-tack tape).

Regardless, the robust performance of these first assays suggests that the strategy may eventually

be useful for a wide range of applications, such as immunoassays or microarray analysis.

As described, the skin depot strategy is similar to the concept of pre-loaded reagents

stored in microchannels.58,59,211-215,219 This is a powerful approach for technology translation, as it

spares the end user from having to procure the reagents and interface them with the experimental

setup. Unlike these previous methods, in which devices are typically disposed after use, in the

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skin depot strategy, the fundamental device architecture can be re-used for any number of assays.

Additionally, because the reagents (and the resulting products) are not enclosed in channels, they

are in an intrinsically convenient format for analysis. For example, in this work, the format was

convenient for MALDI-MS detection, but we speculate that a wide range of detectors could be

employed in the future, such as optical readers or acoustic sensors.

5.3.4 Skin Depot Stability

To be useful for practical applications, pre-loaded reagents must be able to retain their

activity during storage. To evaluate the shelf-life of skin depots, we implemented a quantitative

protein digest assay. The reporter in this assay, quenched BODIPY-labeled casein, has low

fluorescence when intact, but becomes highly fluorescent when digested. In skin depot assays, a

droplet containing the reporter was driven to a pre-loaded spot of trypsin, and after incubation

the fluorescent signal in the droplet was measured in a plate reader (as described

previously90,105,220). In preliminary experiments with freshly prepared skins, it was determined

that at the concentrations used, the reaction was complete within 30 minutes. An internal

standard (IS), rhodamine B, was used to correct for alignment errors, evaporation effects, and

instrument drift over time.

In shelf-life experiments, skins were stored for different periods of time (1, 2, 3, 10, 20,

or 30 days) at -20°C or -80°C. In each experiment, after thawing the skin, positioning it on the

device, driving the droplet to the trypsin, and incubating for 30 minutes, the reporter/IS signal

ratio was recorded. At least five different skins were evaluated for each condition. As shown in

Figure 5.6, shelf-life performance was excellent – substrates stored at -80°C retained >75% of

the original activity for periods as long as 30 days. Substrates stored at -20°C retained >50% of

113

the original activity over the same period. The difference might simply be the result of different

average storage temperature, or might reflect the fact that the -20°C freezer was used in auto-

defrost mode (with regular temperature fluctuations), while the temperature in the -80°C freezer

was constant. Regardless, the performance of these substrates was very encouraging for a first

test, and we anticipate that the shelf-life might be extended in the future by adjusting the enzyme

suspension buffer pH or ionic strength or by adding stabilizers (e.g., disaccharides used widely

for dry protein preservation221).

100

80

60

40

20

0

% A

ctiv

ity

Day 1 Day 2 Day 3 Day 10 Day 20 Day 30

-80C

-20C

Figure 5.6: Skin depot stability assay. The fluorescence of protease substrate (BODIPY-casein) and an internal standard were evaluated after storing skin depots for 1, 2, 3, 10, 20, and 30 days. The substrates were stored at -20°C (red) or -80°C (blue). The mean response and S.D. were calculated for each condition from 5 replicate skins.

114

5.4 Conclusion

We have developed a new strategy for DMF, relying on removable plastic coverings, or

skins. This strategy facilitates virtually un-limited re-use of devices without concern for cross-

contamination, as well as enabling rapid access to pre-loaded reagents. As a first solution to the

world-to-chip interface problem for the DMF format, this innovation has the potential to make

this technology attractive for a wide range of applications.

115

Concluding Remarks and Future Potentials

The trend towards laboratory miniaturization is leading to the necessity of robust and

reliable interfaces between MS and microfluidics. Although general tools for wide-spread use are

not yet available, it is clear that technologies are becoming more effective and reproducible.

Here, I summarize my contributions in this area and propose suggestions for future work.

Microfluidic-ESI-MS Interfaces (Chapter 2, 3, and 4)

Chapter 2 presented a method for fabricating a nanospray ionization tip for interfacing

microfluidic devices to MS. The construction of the chip was simple, and the method fell under

"spray from chip" approach; however, in contrast to previous methods reported, eluent spreading

was significantly reduced by (1) the use of thin substrates (total thickness of assembled device

~300µm), (2) alignment of the tip at corner of the substrate (thus benefitting from a naturally

tapered shape), and (3) the use of a relatively hydrophobic substrate (Parylene-C). The

performance of this device was comparable to commercially available pulled glass emitter and

Agilent HPLC ChipTM. In this preliminary work, the dimension of the tip constructed was 20 µm

deep by 60 µm wide. Since Schilling et al.118 reported that reducing the diameter of the nozzle

increases the sensitivity, stability, and durability of the spray even for 100% aqueous solution, a

potential future improvement of this method would be building devices with smaller tip openings

to fully leverage the advantages of the lab-on-a-chip concept, devices should be built with more

complex channel geometries to affect sample injections and separations. In addition, spray

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stability will be evaluated for a longer period of time, the effect of changing the composition of

the solvent (i.e., gradient) will be monitored.

Chapter 3 presented a new method for sample processing and analysis of markers of

amino acids in dried blood samples using an integrated microfluidic device. The integrated

device built on the parylene-glass tips described in Chapter 2. In this work, sample droplets are

manipulated on a top surface, and are subsequently transferred to microchannels on the bottom

of the device through a vertical hole. The spray was generated at the corner of the device by

applying a high voltage to the counter electrode (closed DMF format). This method has the

potential to contribute to a new paradigm of fast, inexpensive screening of analytes in newborn

blood and for other applications. In future work, I propose that it would be useful to explore

building a large multiplex DMF platform capable of analyzing multiple samples (e.g. 12

capillary emitters) in parallel. In addition, given the resurgence in popularity of dried blood spot

analysis,159 I propose that it would be interesting to evaluate many different applications,

including pharmaceutical/metabolite screening and pen-side analysis using the new integrated

DMF platform.

Chapter 4 presented a simpler and more direct method for coupling DMF with ESI-MS

as compare to previous Chapter. The method involves decoupling the spray from DMF processes

via an interface formed by sandwiching a pulled glass capillary emitter between the two DMF

substrates. In addition, the new method incorporates a feedback control system enabling facile,

high-fidelity droplet movement without any manual intervention. As proof of concept, on-chip

processing and analysis of succinylacetone from dried blood spots was performed. This is still a

117

work in progress, and the next step will be validating the method by quantifying succinylacetone

from several quality control samples provided from Children’s Hospital of Eastern Ontario, and

then compare the results obtained using this method with those obtained using conventional

techniques.

Microfluidic-MALDI-MS Interface (Chapter 5)

Chapter 5 presented a new strategy for DMF, in which a removable plastic “skin” was

used to (a) eliminate cross-contamination, (b) bridge the world-to-chip interface, and (c)

interface with MALDI-MS. This method allowed many biochemical reactions to be performed

using DMF techniques on a single device, followed by off-line MALDI-MS analysis without

worrying about sample cross-talk because of non-specific adsorption of analytes on surfaces. In

addition, it served as a useful medium for reagent introduction and storage onto DMF devices by

preloading analytes onto the skin for future uses. The lifetime of enzymatic substrates stored on

skins could be extended via addition of preservatives (e.g., disaccharides), and the stability of

enzymes will be evaluated against a standard control (e.g., liquid enzyme aliquots stored at -80

ºC) over a month period. While initial proof-of-principle experiments comprise a small number

of reagents, I speculate that in the future, a microarray spotter could be used to form skins

appropriate for high-throughput analysis.

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Appendix A Digital Microfluidic Method for in situ Formation of Porous Polymer Monoliths with Application to

Solid Phase Extraction

We introduce the marriage of two technologies: digital microfluidics (DMF), a technique

in which droplets are manipulated by application of electrostatic forces on an array of electrodes

coated by an insulator, and porous polymer monoliths (PPMs), a class of materials that is popular

for use for solid phase extraction and chromatography. In this work, circular PPM discs were

formed in situ by dispensing and manipulating droplets of monomer solutions to designated spots

on a DMF device followed by UV-initiated polymerization. We used PPM discs formed in this

manner to develop a digital microfluidic solid phase extraction (DMF-SPE) method, in which

PPM discs are activated and equilibrated, samples are loaded, PPM discs are washed, and the

samples are eluted, all using microliter droplets of samples and reagents. The new method has

comparable extraction efficiency (93%) to pipette-based ZipTips, and is compatible with

preparative sample extraction and recovery for on-chip desalting, removal of surfactants, and

preconcentration. We anticipate that DMF-SPE may be useful for a wide range of applications

requiring preparative sample cleanup and concentration.

119

A1.1 Introduction

In the past two decades, there has been great interest in the development of micro total

analysis systems (µTAS) that integrate sample delivery, separation and detection on miniaturized

devices.222 However, a major challenge for such methods is sample complexity -- many samples

require processing prior to analysis to purify and extract the desirable analytes. Among the

various sample processing techniques, the most prominent method is solid phase extraction

(SPE), which exploits interactions between a liquid sample and a solid stationary phase material.

In a typical SPE, the sample is retained by a stationary phase and is subsequently eluted in a

more concentrated, purified form.

Microfluidic SPE techniques reported previously have taken many different forms, using

solid-phase materials formed on channel walls,223,224 packed bed of beads,225-232 porous

membranes,233-236 and porous polymer monoliths (PPMs).69,237,238 The latter technique, relying on

PPMs, is particularly attractive because of the capacity to easily form the stationary phase in

microchannels (i.e., by filling channels with a solution of monomers followed by UV- or heat-

curing to form polymer monoliths). Despite these promising developments, the microfluidic SPE

methods reported previously are not a perfect match for preparative applications, as samples

handled in microchannels are often difficult to recover.239 Ideally, a method could be developed

combining the advantages of µTAS (i.e., integration of many automated steps on a single

platform) with the capacity to perform preparative sample cleanup, such that samples could be

recovered and used for various applications.

An alternative miniaturized fluid handling format to microchannels is digital

microfluidics (DMF), a technique in which discrete fluidic droplets are manipulated by

120

electrostatic forces on an array of electrodes coated with an insulating dielectric.75,76,78 While

DMF shares many characteristics with microchannels, DMF is a distinct paradigm from

microchannels. In DMF, droplets can be dispensed, merged, mixed and split independently from

each other, making this technique appropriate for carrying out multistep reactions. Moreover,

DMF is particularly well-suited for preparative processes, as samples with volumes as large as

milliliters240 can be manipulated. We have used DMF previously for many preparative

applications in the past, including processing of samples containing proteins91,92 and hormones241

followed by analysis off-line by mass spectrometry. Here, we report a new digital microfluidic

method to form PPM discs in situ, with application to on-chip SPE. This is the first example of

combining PPMs with digital microfluidics, and we anticipate that this combination may be

useful for a wide range of applications requiring preparative sample cleanup and concentration.

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A1.2 Materials and Methods

A1.2.1 Reagents and Materials

Unless otherwise stated, all chemicals were obtained from Sigma-Aldrich (Oakville,

ON) and were used without further modification. All buffers were formed using deionized water

that had a resistivity of 18 MΩ·cm at 25°C, filtered with nylon syringe filters from Millipore

(Billerica, MA, 0.2 µm pore diameter) and sonicated (5 min) prior to use. Ethanol (95%) and

NaOH were purchased from ACP Chemicals (Montreal, QC). NaCl was purchased from

Mallinckrodt Baker (Phillipsburg, NJ). Fluorescein was purchased from Life Technologies

(Burlington, ON). Cleanroom reagents and supplies included Parylene C dimer from Specialty

Coating Systems (Indianapolis, IN) and Teflon-AF from DuPont (Wilmington, DE). C18

ZipTips were purchased from Millipore.

All solutions used on DMF devices were used within three days of preparation. A C12

casting solution was prepared by mixing 279 µL of butyl acrylate, 150 µL of 1,3-butanediol

diacrylate, 69 µL of lauryl acrylate, 2.5 mg of 2,2-dimethoxy-2-phenylacetophenone, and 1 mL

of porogen comprising a 4:1:1 ratio of acetonitrile, 95% ethanol, and 5 mM phosphate buffer at

pH 6.8. For digital microfluidic solid phase extraction experiments, activation solvent was 0.1%

(v/v) formic acid in acetonitrile, equilibration and wash solvents were 0.5% (v/v) formic acid in

deionized water, and elution solvent was 500 mM sodium borate in deionized water at pH 9.0

(for elution of fluorescein) or 0.1% (v/v) formic acid in acetonitrile (for elution of peptides). A

fluorescamine-labelled peptide standard was prepared using methods similar to those described

previously.242 Briefly, a stock solution was formed by mixing 100 µL fluorescamine (3 mg/mL

in acetone), 10 µL of angiotensin IV (10 mM in 10 mM borate buffer at pH 9), and 890 µL of

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acetone. The reaction mixture was allowed to incubate for at least 2 hours before being diluted

10x with 0.1% formic acid for analysis.

A1.2.2 Device Fabrication and Operation

Digital microfluidic devices were fabricated in the University of Toronto Emerging

Communications Technology Institute (ECTI) cleanroom facility, using a transparent photomask

printed at Pacific Arts and Design (Markham, ON). Glass devices bearing patterned chromium

electrodes were formed by photolithography and etching as described previously,98 and were

coated with 7 µm of Parylene-C and 50 nm of Teflon-AF. Parylene-C was applied using a vapor

deposition instrument (Specialty Coating Systems), and Teflon-AF was spin-coated (1% wt/wt in

Fluorinert FC-40, 1000 rpm, 30 s) followed by post-baking on a hot-plate (160ºC, 10 min). The

polymer coatings were removed from contact pads by gentle scraping with a scalpel to facilitate

electrical contact for droplet actuation. In addition to patterned devices, unpatterned indium tin

oxide (ITO) coated glass substrates (Delta Technologies Ltd, Stillwater, MN) were coated with

Teflon-AF (50 nm, as above).

Two device designs were used, shown in Figures A1.1a and A1.5a. The former featured

an array of eighty-eight actuation electrodes (2.2 x 2.2 mm ea.) connected to ten reservoir

electrodes (5 x 5 mm ea.), with inter-electrode gaps of 40 µm. The latter design featured 12 large

actuation electrodes (7.5 mm x 7.5 mm ea.) for moving sample and solvent droplets, and 19

small actuation electrodes (2.2 mm x 2.2 mm ea.) for moving monolith casting solution and

elution solvent droplets. Devices were assembled with an unpatterned ITO–glass top plate and a

patterned bottom plate separated by a spacer formed from three pieces of double-sided tape (total

123

spacer thickness 270 µm), such that "unit" droplets (i.e., the droplet size covering a single

electrode) were 1 µL for 2.2 x 2.2 mm electrodes and 12 µL for 7.5 mm x 7.5 mm electrodes.

To actuate droplets, driving potentials (220-300 Vpp) were generated by amplifying the output of

a function generator (Agilent Technologies, Santa Clara, CA) operating at 18 kHz. As described

elsewhere,91,98 droplets were sandwiched between a patterned bottom plate and an unpatterned

(Teflon-ITO-glass) top plate and actuated by applying driving potentials between the top

electrode (ground) and sequential electrodes on the bottom plate via the exposed contact pads.

Droplet actuation was monitored and recorded by a CCD camera mounted on a lens. For the

device shown in Figure A1.1a, 1 µL unit droplets were dispensed from larger volumes in

reservoirs by actuating three adjacent electrodes in series as described previously.243 On the same

device design, larger 5 µL droplets were dispensed from reservoirs and actuated by applying

potentials to multiple electrodes simultaneously.

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Activationsolvent

Wash/Equilibrationsolvent

Casting solutionSample

Elution solvent

Waste/CollectorUV polymerizedPPM disc

Top Plate

Bottom Plate

PPMdisc

Air ~

Electrode HydrophobicDielectric Glass

1casting solution

2

PPM disc

5 µm

1 mm

a) DMF Device

b) Formation of PPM disc

c) Images of PPM disc

Figure A1.1: DMF device design and PPM disc. (a) Schematic of the device, which features reservoirs dedicated to five different reagents, including elution solvent, sample, monomer, wash solvent, and activation solvent. The reservoirs are connected to an array of eighty-eight actuation electrodes. (b) Frames from a movie illustrating the formation of a PPM disc. In frame 1, a 1 µL droplet of casting solution was dispensed and driven to the middle of the device. The PPM disc was formed (frame 2) as a result of exposure to UV irradiation (100 W, 365 nm, 5 min). (c) A brightfield microscope image of a PPM disc formed on chip (left) and a high-resolution SEM image of the interior of a PPM disc (right). Scale bars are 1 mm and 5 µm, respectively.

125

A1.2.3 PPM Formation and Characterization

Porous polymer monoliths (PPMs) were prepared via on-chip photopolymerization of a

C12 casting solution droplet manipulated by DMF. As illustrated in Figure A1.1b, each PPM

was formed by dispensing a 1 µL droplet of casting solution onto the array of actuation

electrodes and translating it to a central position where it was polymerized by exposure to UV

radiation (100 W, 365 nm, 5 min). As shown, the resulting PPM took the shape of a circular disc

with radius ~1 mm and height of 270 µm. Two additional types of PPMs were formed, in (a) a

glass pipette, and (b) in a "dummy substrate." The former (in pipette) was used to evaluate the

quality of the casting solution. These PPMs were formed by aspirating ~50 µL of casting

solution into the glass pipette and exposing to UV radiation as described above. The latter (in

dummy substrates) was required for N2 adsorption experiments because the method used (see

below) was not sensitive enough evaluate the tiny PPM discs formed in digital microfluidic

devices. These discs were formed by polymerizing 15 µL droplets sandwiched between pieces of

unpatterned glass coated with 50 nm Teflon-AF (formed as above) with 450 µm spacing between

them. PPMs formed in this manner had approximate diameters of 5 mm and masses of 3.1 mg.

The surface morphologies and surface areas of PPMs were characterized by scanning

electron microscopy (SEM) and N2 adsorption, respectively. Surface morphologies of PPMs

formed on-chip and in glass pipettes were observed using a Hitachi S-5200 SEM (Hitachi,

Mississauga, ON); prior to analysis, specimens were carbon-coated to prevent charging. Surface

areas of PPMs formed in dummy substrates were determined using an Autosorb-1 instrument

from Quantachrome Corporation (Boynton Beach, Florida). The samples were outgassed at 50°C

for 4 h under vacuum, and the specific surface area (SSA) was obtained from N2 (purity

126

99.999%) adsorption at 77 K using Brunauer, Emmett, and Teller (BET) theory.244 For the

calculation of the BET SSA, the relative pressure range was assumed to be 0.05 – 0.1 mbar. At

least three different PPMs were evaluated for all characterization experiments.

A1.2.4 Standard DMF-SPE Process

All digital microfluidic solid phase extraction (DMF-SPE) experiments were carried out

using a variation on the following four-step procedure. Step 1: Activation/Equilibration. A PPM

disc was activated by dispensing and driving a 5 µL droplet of activation solvent onto the

monolith. The droplet was actuated back and forth across the PPM disc five times and then was

allowed to incubate on the disc for 1 minute before it was driven away and collected in the waste

reservoir. An identical process (dispense, actuate, incubate, move to waste) was then

implemented for a 5 µL droplet of equilibration solvent. Step 2: Sample Loading. A 1 µL sample

droplet was dispensed and actuated to the PPM and was actuated back and forth across the PPM

disc five times before being incubated for 2, 4, or 8 minutes, and then the droplet was moved to

waste. Step 3: Monolith Washing. Droplets of wash solvent (2 x 5 µL) were dispensed, actuated

onto the monolith (and back and forth across the PPM disc five times) and then moved to waste.

Step 4: Elution. One, two or three 5 µL droplets of elution solvent were dispensed, actuated onto

the monolith (and back and forth across the PPM disc five times) and then moved to a sample

collection reservoir.

A1.2.5 DMF-SPE Process Optimization

Fluorescein (5 µM in 0.5% v/v aqueous formic acid) was used as a model analyte to

optimize the DMF-SPE process on the device shown in Figure A1.1a for (i) sample loading time,

and (ii) elution volume. For the former (optimization of sample loading time), a truncated DMF-

127

SPE method was performed incorporating only steps 1 and 2 from above. After incubating the

sample droplet on the PPM for 2, 4, or 8 minutes, the sample droplet was moved to a fresh spot

on the device and dried by heating on a hot plate (50°C, ~2 min). The precipitate was manually

resolubilized in 50 µL of borate buffer (500 mM, pH 9) and transferred to a 384-well plate for

fluorescence measurement (λex: 480 nm, λem: 520 nm) using a PHERAstar plate reader (BMG

Labtech, Durham, NC). For the latter (optimization of elution volume), the full four-step DMF-

SPE process was used (with a 2 minute incubation in step 2). In step 4, the samples were eluted

in 1, 2, or 3 droplets of elution solvent, which were pooled and moved to a fresh spot on the

device, and dried, resolubilized, and interrogated for fluorescence intensity as described above.

For both optimizations (i and ii), the fluorescence intensities of the experimental samples were

compared to the fluorescence intensities of control experiments in which 1 µL fluorescein

droplets were driven across the device without any exposure to a PPM and then dried and

resolubilized as above. All experimental and control experiments were conducted in triplicate

using three different PPMs/devices.

A1.2.6 SPE Extraction Efficiency

A fluorescamine-labeled peptide (10 µM angiotensin IV in 0.1% formic acid) was used

as a model analyte to evaluate the extraction efficiency of DMF-SPE on the device shown in

Figure 1a and by commercially available ZipTips. For the former (DMF-SPE), the labeled

peptide sample was extracted using the four-step procedure described above with a 2-minute

incubation in step 2 and with two droplets of elution solvent in step 4. The eluted droplets were

pooled and moved to a fresh spot on the device, and dried, resolubilized in 50 µL of elution

solvent, and interrogated for fluorescence intensity using the plate reader (with λex: 390 nm, λem:

128

460 nm) as described for process optimization, above. For the latter (conventional SPE), 5 µL

aliquots of labelled peptide sample were extracted on C18 ZipTips as per the manufacturer’s

instructions. The extracted samples were dried and dissolved in 50 µL of elution solvent for

analysis using the plate reader. For both cases (DMF-SPE and ZipTip), the fluorescence

intensities of the experimental samples were compared to the fluorescence intensities of control

samples (1 µL for DMF-SPE, 5 µL for ZipTip) which were not extracted, but dried and

resolubilized as above. All experimental and control experiments were conducted in triplicate

using three different PPM/devices or ZipTips.

A1.2.7 DMF-SPE for Preparative Sample Cleanup

DMF-SPE was validated for use for preparative sample cleanup by extraction of two

solutions of angiotensin II (Ang II, 1 µM) on the device shown in Figure A1.1a: (1) in a solution

containing 100 mM sodium chloride, and (2) in a solution containing 0.05% w/v Pluronic F68.

For both experiments, 1 µL sample droplets were extracted using the four-step procedure

described above with a 2-minute incubation in step 2 and with two droplets of elution solvent in

step 4. The eluted droplets were pooled and moved to a fresh spot on the device, and dried at

room temperature. The eluate was manually resolubilized in 50 µL of 50% acetonitrile

containing 0.1% formic acid, and analysed via nanoESI-MS (LTQ Finnigan, Thermo Electron

Corp., FL). The applied spray voltage was varied between 1.7-2.0 kV and the flow rate of the

syringe pump and capillary temperature were kept constant at 0.5 µL/min and 200°C,

respectively. As a control, 1 µL aliquots of the same samples (without desalting or removal of

surfactant) were dried, resolubilized and analyzed by nanoESI-MS for comparison. Each

experiment and control was performed in triplicate using separate devices/PPMs.

129

A1.2.8 DMF-SPE for Sample Concentration

The device shown in Figure 5a was designed to evaluate the potential of DMF-SPE for

sample concentration. A variation of the four-step procedure described above was implemented

with fluorescein as the sample (5 µM in 0.5% v/v aqueous formic acid). In steps 1-3, the

volumes of activation, equilibration, sample, and wash droplets were 12 µL (instead of 1 or 5

µL). Step 4 was similar to the procedure outlined above but used a single 1 µL elution droplet.

After the extraction was complete, the eluted droplet was pipetted off the device and diluted in

49 µL of elution solvent. The fluorescence intensity was measured using a plate reader as above

and compared with that of a control sample comprising 1 µL of fluorescein sample, not

extracted, diluted with 49 µL of borate buffer (500 mM, pH 9).

130

A1.3 Results and Discussion

A1.3.1 PPM Formation and Characterization

We report here the first combination of digital microfluidics and porous polymer

monoliths (PPMs). PPMs have been used previously in enclosed microchannels69,122,237,245-247 and

capillaries248-250 by polymerizing a casting solution containing monomers that is surrounded on

all sides by the walls of the channel or capillary. Monoliths formed in this manner take the shape

of the channel or capillary containing the casting solution. Here, we introduce a new format for

PPMs, in which droplets of casting solution are dispensed onto an array of electrodes where they

are manipulated into position and then polymerized. This format confers some advantages: the

PPMs can be formed in any desired location on the device, multiple PPMs may be formed in

parallel, and the PPMs are open on all sides (in the plane of the device), which facilitates a wide

range of solid-liquid interactions.

A schematic of a device is shown in Figure A1.1a, and pictures depicting the formation of

a monolith are shown in Figure A1.1b. As shown, PPMs formed in this manner take the shape of

a circular disc which is slightly smaller than the droplet from which it was formed (e.g., a 1 µL

droplet with ~2.2 mm dia. forms a disc with ~2 mm dia.). The size of the discs is fairly

reproducible -- for example, when five discs were formed on five different devices, the

coefficient of variation in diameter was 10%. Magnified images of PPM discs formed on DMF

devices are shown in Figure A1.1c. The discs have a skin-like outer layer and an interior with the

characteristic popcorn-like structure to PPMs formed in conventional formats. The average

surface area of PPM discs formed on substrates approximating DMF devices was found to be 3.7

131

± 0.2 m2/g by N2 adsorption, a value that is similar to what has been reported for PPMs formed

in capillaries.251

To our knowledge, this paper describes the first combination of digital microfluidics with

three-dimensional structures positioned on an array of DMF electrodes. As we began this work,

we hypothesized that there might be two potential pitfalls. First, because droplets manipulated by

DMF are known to pin (i.e., get stuck) on heterogeneous regions on device surfaces,208 we

hypothesized that droplets might experience similar problems when they encountered PPM discs.

In fact, this was observed for cases in which the droplets were smaller than PPM discs -- in such

cases, droplets were observed to become stuck on the PPMs resulting in no further movement.

To circumvent this problem, we used sample or solvent droplets that were larger than the PPMs,

such that a small fraction of the droplet could stick to the monolith, while a large fraction of the

droplet could move away. We have observed similar phenomena, dubbed "passive dispensing,"

for the process of translating a droplet across a patterned hydrophilic region for cell culture.97 For

the 2.2 mm dia./1 µL droplets and ~2 mm. dia. PPM discs used here, we estimate that when a

droplet is driven onto and off of a monolith disc, the volume remaining on the monolith is less

than 10% of the original droplet volume. Second, there was concern that PPMs (which, in this

case, are not covalently bound to the device surfaces and can be removed/replaced using

tweezers when the device is disassembled) might become mobile when they encounter moving

droplets. Interestingly, this phenomenon was not observed during the course of hundreds of

experiments. We hypothesize that the friction forces between the PPM discs and the device

surfaces are larger than the forces associated with moving droplets.

132

A1.3.2 DMF-SPE

After developing means to form PPM discs in situ on digital microfluidic devices, a four-

step procedure was developed to implement digital microfluidic solid phase extraction (DMF-

SPE), which is depicted in Figure A1.2. First, the PPM was activated in a non-polar solvent and

then equilibrated in a polar solvent (Figure A1.2a). Second, a small sample droplet was loaded

(Figure A1.2b), and third, any unbound sample was washed away by a large droplet of wash

solvent (Figure A1.2c). Fourth, a droplet of elution solvent was used to extract the target analyte

from the PPM disc (Figure A1.2d).

A fluorescence method was developed to optimize the DMF-SPE procedure for loading

time and elution volume, using fluorescein as a model substrate. For the former (loading time),

fluorescein droplets were allowed to incubate on PPM discs for 2, 4 or 8 minutes before

collecting the sample droplet and measuring the remaining analyte as a function of the

fluorescence intensity. As shown in Figure A1.3a, after incubating for 2 minutes, less than 20%

of the fluorescence (relative to the control) remained in the sample droplet. A t-test revealed no

significant differences between 2 or 4 minutes of incubation (p = 0.30), so a 2-minute incubation

period was used for all subsequent experiments. For the latter (elution volume), fluorescein was

loaded onto PPM discs, rinsed, and then eluted in one, two or three 5 µL droplets of elution

buffer. As shown in Figure 3b, 2 elution droplets extracted over 90% of bound sample from the

PPMs. A t-test revealed no significant difference (p = 0.25) between 2 droplets and 3 droplets, so

2 elution droplets were used for all subsequent experiments.

133

(a) Activation/Equilibration of PPM disc

PPM Disc

solventActivation/Equilibration

1Incubate

2Move to waste

3

(b) Sample Loading

2Incubate

3

Move to waste

1

Sample droplet

(c) Washing

1

Wash solvent

2Incubate

3

Move to waste

(d) Elution

1

Elution solvent

2Incubate

3

Move to collector

Figure A1.2: DMF-SPE protocol. Frames from a movie (left-to-right) depicting the four-step digital microfluidic solid phase extraction procedure. In (a), a PPM disc is activated by dispensing a 5 µL droplet (frame 1) of activation solvent and driving it onto the monolith. After incubation (frame 2), the droplet is moved away to the waste reservoir (frame 3). Similar steps are repeated for PPM equilibration (not shown), followed by sample loading (b), PPM washing (c), and sample elution (d).

The optimized DMF-SPE method was validated by comparing its extraction efficiency to

that of the commercially available ZipTip method. Fluorescamine-labelled angiotensin IV was

used as a model substrate, and after comparison with controls, the DMF-SPE method had an

extraction efficiency of 93 ± 14% and the ZipTip had an extraction efficiency of 92 ± 5%. This is

134

a remarkable similarity given that the DMF-SPE monolith was formed with C12 monomers, and

the ZipTip possessed C18 functionality. We propose that in the future, methods for forming

PPMs with C1869 or other functional groups237,248,249 might be adapted for use in DMF. In

addition, future methods will be combined with automated feedback-controlled dispensing252 to

control and reduce the variability of dispensed droplet size, which we speculate will improve the

precision of the DMF-SPE technique.

100

80

60

40

20

00 2 4 8

Loading time (min)

Loading Time Optimization

Rel

ativ

e F

luor

esce

nce

Control

# of elution droplets

Rel

ativ

e F

luor

esce

nce 160

120

80

40

01 2 3

Elution Optimization

a) b)

Figure A1.3: DMF-SPE optimization. Loading time (a) was optimized by evaluating the fluorescence of droplets that had incubated 2, 4, or 8 minutes on a PPM disc relative to a control sample. Elution volume (b) was optimized by evaluating the number of elution droplets required to extract a fluorescent analyte from a PPM disc

A1.3.3 DMF-SPE for Preparative Analysis

To validate DMF-SPE for preparative processes, two applications were implemented on-

chip followed by analysis by mass spectrometry: sample desalting and removal of surfactant. For

the former (desalting), a solution containing 1 µM angiotensin II and 100 mM NaCl was

evaluated. As shown in Figure A1.4a, the non-extracted sample had no peak at m/z = 1047 (the

[M+H]+ peak for angiotensin II) because of ion suppression by the salt matrix, but after

135

desalting, a strong signal at m/z = 1047 was observed. For the latter (removal of a surfactant), a

solution containing 1 µM angiotensin II and 0.05% Pluronic F68 was evaluated. This case is

particularly important for digital microfluidics, as we90-92,97,98,101,105 and others253,254 have used

various Pluronics as solution additives to prevent adsorption of peptides and proteins onto DMF

device surfaces. Unfortunately, Pluronics, like other surfactants, are known to be strong ion

suppressors for MS,9,10 making the ability to remove the additives prior to analysis critical for

obtaining useful information. As shown in Figure A1.4b, the non-extracted sample had no peak

at m/z = 1047, whereas the extracted sample (Figure A1.4b) had a strong signal.

1500135012001050900

Extracted

140012001000

Non-extracted

[M + H]+

[M + Na]+

[M + K]+

1500135012001050900

140012001000

Extracted Non-extracted

[M + H]+

a) NaCl b) Pluronic F68

m/z m/z

Figure A1.4: Preparative DMF-SPE. NanoESI mass spectra generated from solutions of angiotensin II (MW 1046) containing (a) 100 mM NaCl, or (b) Pluronic F68 (0.05% w/v). The spectra in the main panels were generated from solutions that had been extracted, and the spectra in the insets were generated from solutions that had not been extracted. The data shown in Figure A1.4 are superficially similar to data obtained with methods

demonstrated previously for sample preparation for MALDI-MS.93,94 In the previous work,

sample droplets were allowed to dry on flat DMF device surfaces, after which droplets of water

were driven over the spots to rinse them. The dried samples were found to be substantially

purified. This previous method93,94 represented an important milestone for DMF, but the new

136

method reported here is likely superior for several reasons. First, the three-dimensional PPM

discs reported here have a larger surface area than a flat surface of equal size, which suggests

that the analyte loading capacity for the new method is higher. Second, in the new method,

analytes are eluted from the PPM disc and can be delivered to any desired location, whereas in

the previous method, the sample is permanently bound to a particular spot on the device surface.

Third, PPMs can be formed with a wide variety of chemical functional groups,69,237,248,249 while

the previous method is limited to the device surface material (Teflon-AF).

A1.3.4 DMF-SPE for Sample Concentration

A critical application for SPE is sample concentration, and a new device, shown in Figure

A1.5a, was designed to probe this application for DMF-SPE. This device featured a combination

of large electrodes used to deliver 12 µL sample droplets, and small electrodes which were used

to deliver 1 µL elution solvent droplets. The process is illustrated in Figure A1.5b, in which a

large sample droplet is loaded onto a PPM disc followed by elution in a much smaller droplet.

Using this design, a concentration factor of ~9 for sample droplets of fluorescein was achieved as

shown in Figure A1.5c.

The data and method depicted in Figure A1.5 are preliminary. The concentration factor of

~9 was smaller than the expected value of 12. We speculate that the results presented here will

be improved in future work, when PPM disc size and shape and electrode layout are optimized.

Regardless, this proof-of-principle data fits comfortably in the range of concentration factors for

ZipTips (i.e., 2.5-10), and suggests that DMF-SPE may a useful new tool for of applications that

are commonly executed using the more conventional technique.

137

a

Activationsolvent

Wash/Equilibrationsolvent

Sample

Elution solvent

Waste/Collector

UV polymerizedPPM disc

Monomer

Sample (12 µL) Eluent (1 µL)

Concentrated Extract

Control

Rel

ativ

e F

luor

esce

nce

10

8

6

4

2

0

a)

b)

c)

Figure A1.5: DMF-SPE for concentration. (a) Schematic of device featuring large electrodes (7.5 mm x 7.5 mm) for sample, activation, and washing solvents and small electrodes (2.2 mm x 2.2 mm) for elution solvent and monomer casting solution. (b) Frames from a movie illustrating the process of concentrating a sample on-chip using a PPM disc. In frame 1, a 12 µL sample droplet was actuated onto an activated PPM disc. In frame 2, the disc was washed and a 1 µL droplet of elution buffer was driven onto it to extract the sample. (c) Bar graph demonstrating a concentration factor of ~9 in extracted samples versus controls.

138

A1.4 Conclusion

We have successfully combined two technologies, digital microfluidics (DMF) and

porous polymer monoliths (PPMs). In this work, PPM discs were formed in situ on DMF devices

by dispensing droplets of monomer solutions onto an array of electrodes followed by UV-

initiated polymerization. The PPM discs were used for preparative solid phase extraction (SPE)

with all fluidic handling steps carried out by DMF. The new method had comparable extraction

efficiencies to that of commercially available ZipTips, and was compatible with straightforward

sample extraction and recovery. We anticipate that this technique may be useful for a wide range

of applications requiring preparative sample cleanup and concentration.

139

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