development of a miniaturized chromatographic analytical device … · requiring high-throughput...

Development of a miniaturized chromatographic analytical

device for food safety applications

Catarina Isabel Pereira Bombaça

Thesis to obtain the Master of Science Degree in

Biotechnology

Supervisors: Prof. Dr. João Pedro Estrela Conde and Dra. Ana Margarida Fortes

Examination Committee

Chairperson: Prof. Dr. Luís Joaquim Pina da Fonseca

Supervisor: Prof. Dr. João Pedro Estrela Conde

Members of the Committee: Dr. Ana Margarida Nunes da Mata Pires de Azevedo

November 2017

iii

Acknowledgments

In first place, I would like to thank for the support, motivation and mentorship given by my

supervisors Prof. Dr. João Pedro Conde and Dra. Ana Margarida Fortes. I really would like to emphasis

my gratitude in allowing me to work on this project. I would also like to thank Dra. Virginia Chu for the

support given throughout my year in INESC-MN.

I would like to thank my colleagues from INESC-MN for the amazing experience that has been

working together this last year. I will never forget the supportive environment provided during work, with

a special thanks to Inês and Ruben who were dedicated and amazing mentors in every phase of this

project. Not only them, but everyone else who contributed in some way to help me grow during this

project, and for all the fun times and cheerful spirit in the lab!

I cannot forget my friends from NOVA with whom I shared the most important moments and

experiences during this academic journey, particularly Gonçalo, my favourite Azorean and partner on

the many adventures we shared in Czech Republic and the rest of Europe. To my friends from Board of

European Students of Technology, for all the hard-working days and nights when we did not even slept

to pursue this dream we shared. A special thanks to Palmeiro, Tomás and Sara who were very

supportive in all projects I pursued, including this one. And, of course, a special thanks to Ľuboš and

Sachin for motivating me to work, while I was doing my thesis, when it was the hardest to keep

everything on track: I hope we have the opportunity to meet again as we did in Latvia.

To my childhood friends, Magda and Margarida, for proving that despite the absence, our friendship

prevails and even becomes stronger.

To my parents, for understanding why I was never home and for always being happy for my

achievements, even when they did not quite understand what it meant. Especially, I want to thank my

mom who proved this year to be the strongest woman I ever known and for teaching me that a disease

cannot define who you are.

Finally, to Eurico, for being my favourite person in the world and for supporting me in all the good

and bad times, for the friendship and love, and for dreaming with me all the adventures we are yet to

live throughout this world.

v

Resumo

Existe uma procura cada vez maior em miniaturizar a instrumentação usada em separações

analíticas, exigindo-se respostas que providenciem soluções de elevado rendimento, mas mais

importante ainda, análises rápidas e on-spot. É, portanto, uma prioridade desenvolver um dispositivo

analítico miniaturizado e cromatográfico que atende não só ao objetivo de total integração, mas também

aponta para um processo de fabricação mais simples (economicamente viável), uma operação mais

user-friendly e uma versatilidade para uma ampla variedade de analitos/aplicações.

O desenvolvimento de um dispositivo de microfluídica, consistindo num injetor de geometria cross-

channel com válvulas integradas e atuadas pneumaticamente (que oferecem quase zero dead-volume),

e num canal de 12 cm de comprimento de separação, permitindo o empacotamento de meio

cromatográfico disponível comercialmente, foi alcançado. O canal de separação foi testado para efeitos

de proof-of-concept com a separação de uma mistura de azul de metileno e corante alimentar amarelo,

seguido de regeneração, provando o grande potencial para aplicações em purificação e separações

analíticas. Além disso, a aplicabilidade deste dispositivo para análises de segurança alimentar foi

avaliada usando o canal de separação para cromatografia de fase-reversa, indicando que com

condições cromatográficas futuramente otimizadas e otimização também do dispositivo, a separação e

deteção de toxinas como a aflatoxina B1 (AFB1) e ocratoxina A (OTA) será possível. Finalmente, a

injeção e progressão de um plug de isotiocianato de fluoresceína (FITC), num injetor cross-channel

com válvulas integradas e atuadas pneumaticamente e adaptado ao canal de separação, foi estudada.

Este estudo ofereceu uma avaliação sobre as otimizações futuras ser feitas no dispositivo de

microfluídica desenvolvido.

Palavras-chave:

Microfluídica, cromatografia, separação, ocratoxina A, aflatoxina B1

vii

Abstract

There is an increased demand for smaller and smaller instrumentation for analytical separations,

requiring high-throughput answers to its problems but more importantly, fast and on-spot analyses of

products of interest. Therefore, it is a priority to develop a miniaturized chromatographic analytical device

which not only meets the goal of fully integration but also aims for a simpler fabrication process (and

therefore more economically viable), user-friendly operation and versatility for a broad range of

analytes/applications.

The development of a microfluidic device which comprises a cross-channel injector with integrated

pneumatic actuated valves that offers almost zero dead-volume and a 12 cm-long separation channel

enabling the packing of most commercial available chromatographic beads was achieved. The

separation channel was tested for a proof-of-concept separation that allowed the separation of a mixture

of methylene blue and yellow food colouring and further regeneration of the channel, proving the great

potential for future purification and analytical applications. Moreover, the applicability of this device for

food safety applications was assessed using a reverse-phase chromatographic mode for the separation

channel, indicating that under the optimal chromatographic conditions and further optimization of the

device design, separation and detection of toxins like aflatoxin B1 (AFB1) and ochratoxin A (OTA) is

possible. Finally, a fluorescein isothiocyanate (FITC) plug injection and progression using the cross-

channel injector with pneumatically actuated valves coupled to the separation channel was studied,

which offered an assessment on further optimizations to be done on the microfluidic device developed.

Keywords:

Microfluidics, chromatography, separation, ochratoxin A, aflatoxin B1

ix

Table of Contents

ACKNOWLEDGMENTS ......................................................................................................................... III

RESUMO ................................................................................................................................................. V

PALAVRAS-CHAVE: ............................................................................................................................... V

ABSTRACT ........................................................................................................................................... VII

KEYWORDS: ......................................................................................................................................... VII

TABLE OF CONTENTS ......................................................................................................................... IX

LIST OF TABLES ................................................................................................................................. XIII

LIST OF FIGURES ................................................................................................................................XV

LIST OF ACRONYMS ..........................................................................................................................XIX

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

1.1. MOTIVATION ................................................................................................................................... 1

1.2. CHROMATOGRAPHY ........................................................................................................................ 2

1.2.1. General concepts............................................................................................................. 2

1.2.1.1. Elution .......................................................................................................................... 3

1.2.1.2. Chromatograms ........................................................................................................... 4

1.2.1.3. Retention time .............................................................................................................. 4

1.2.2. Column performance ....................................................................................................... 5

1.2.2.1. The theoretical plate model of chromatography .......................................................... 5

1.2.2.2. The rate theory and the van Deemter equation ........................................................... 6

1.2.3. HPLC ............................................................................................................................... 6

1.2.3.1. Instrumentation ............................................................................................................ 7

1.2.3.2. Chromatographic modes ............................................................................................. 9

1.3. MICROFLUIDICS............................................................................................................................. 11

1.3.1. General concepts........................................................................................................... 11

1.3.2. Microfabrication ................................................................................................................... 13

1.3.2.1. Photolithography ........................................................................................................... 13

1.3.2.2. Soft lithography technologies ....................................................................................... 14

1.3.3. Chromatography-on-chip ..................................................................................................... 16

1.3.3.1 Column design and fabrication considerations ............................................................. 16

1.3.3.2 Injection and sample volume considerations ................................................................ 17

1.3.3.3 Literature examples on chromatography-on-chip .......................................................... 18

1.4. FOOD SAFETY APPLICATIONS ......................................................................................................... 19

2. MATERIALS AND METHODS .......................................................................................................... 21

x

2.1. MICROFLUIDIC DEVICE FABRICATION ............................................................................................... 21

2.1.1. Hard mask fabrication .......................................................................................................... 21

2.1.2. Master mold fabrication ....................................................................................................... 22

2.1.2.1. Silicon substrate cleaning ............................................................................................. 23

2.1.2.2. 20 μm layer ................................................................................................................... 23

2.1.2.3. 35 μm layer ................................................................................................................... 23

2.1.2.4. 50 μm layer ................................................................................................................... 24

2.1.2.5. 100 μm layer ................................................................................................................. 24

2.1.3. Fabrication of PDMS channels ............................................................................................ 25

2.1.3.1. Process for iteration 1, 2 and microcolumns ................................................................ 26

2.1.3.2. Process for iteration 3 and 4A ...................................................................................... 26

2.1.3.3. Process for iteration 4B ................................................................................................ 26

2.2. MICROFLUIDIC DEVICE MANIPULATION ............................................................................................ 27

2.2.1. Separation channel packing .......................................................................................... 27

2.2.1.1. Beads preparation protocols ...................................................................................... 28

2.2.1.2. Packing method ......................................................................................................... 28

2.2.2. Separation and elution studies ...................................................................................... 29

2.2.2.1. Food colouring separation (proof-of-concept) ........................................................... 29

2.2.2.2. OTA and AFB1 microchannel adsorption and elution ............................................... 30

2.2.2.3. OTA and AFB1 cross-channel injection and elution .................................................. 30

2.2.2.4. Plug optimization ....................................................................................................... 31

3. RESULTS AND DISCUSSION ...................................................................................................... 32

3.1. MICROFLUIDIC DEVICE DESIGN CONSIDERATIONS ............................................................................ 32

3.2. FABRICATION PROCESS CONSIDERATIONS ...................................................................................... 37

3.2.1. Valves thickness optimization ......................................................................................... 37

3.2.2. Microfluidic structure sealing optimization ....................................................................... 38

3.3. SEPARATION CHANNEL PACKING .................................................................................................... 39

3.4. FOOD COLOURING SEPARATION (PROOF-OF-CONCEPT) ................................................................... 43

3.5. TOXINS SEPARATION FOR FOOD SAFETY APPLICATIONS ................................................................... 45

3.5.1. Toxins concentration optimization ....................................................................................... 45

3.5.2. OTA and AFB1 elution studies in microchannels ................................................................ 46

3.5.3. OTA and AFB1 cross-channel injection and elution ............................................................ 48

3.6. PLUG PROGRESSION ASSESSMENT AND OPTIMIZATION .................................................................... 51

4. CONCLUSION AND FUTURE PROSPECTS ............................................................................... 55

REFERENCES ...................................................................................................................................... 58

APPENDIX SECTION ........................................................................................................................... 62

A. OPTIMIZED PROCESSES RUNSHEET ................................................................................................... 62

xi

B. SOLVENT EFFECT INFLUENCE ON FLUORESCENCE SIGNAL FOR OTA AND AFB1 ELUTION IN

MICROCHANNELS ................................................................................................................................. 66

xiii

List of Tables

Table 1. HPLC chromatographic modes are summarized with attached examples present in the

literature. ................................................................................................................................................ 10

Table 2. Materials and equipment required for microfluidic device fabrication. .................................... 21

Table 3. Runsheet comprising the sequential steps involved in each mold fabrication process for the

different design iterations, including iterations where two different master molds are necessary (3 and

4). ........................................................................................................................................................... 22

Table 4. Materials and equipment required for microfluidic device manipulation. ................................ 27

Table 5. The different packing solutions required for each beads type. ............................................... 40

xv

List of Figures

Figure 1. Column chromatography. Left - the standard elements of a chromatographic column include

a matrix supported inside a column, generally made of plastic or glass. A protein solution to be separated

can percolate the column (starts on top). Proteins migrate through the column at different rates because

of different interactions established with the matrix. Right - diagram representing the separation of the

chromatography procedure illustrated on the left. The detected signal at the various stages of elution is

shown. Adapted from the literature16,17.................................................................................................... 3

Figure 2. Chromatogram. Left - elution of components A, B and C is represented by plotting the detector

signal intensity in millivolts versus time of elution in minutes. Right - two-component chromatogram

exemplifying two approaches for improving separation: (a) original with overlapping peaks, (b)

improvement by increase in band separation, (c) improvement by decrease in band widths. Adapted

from the literature16,17............................................................................................................................... 4

Figure 3. Chromatogram of a two-component (I and II) mixture. t0 represents the dead time, tR1 and tR2

the retention times of the components I and II, respectively. W1 and W2 are the band widths of each

component and Δt is the difference in elution time between the two components. Adapted from

Thermopedia. .......................................................................................................................................... 5

Figure 4. Diagram of the chromatographic methods. GSC – gas solid chromatography, GC – gas

chromatography, SFC – supercritical fluid chromatography, LC – liquid chromatography, LLC – liquid-

liquid chromatography, MEKC – micellar electrokinetic chromatography, LSC – liquid solid

chromatography, SEC – size exclusion chromatography, IEC – ion exchange chromatography, AC –

affinity chromatography and BPC – bonded phase chromatography. .................................................... 7

Figure 5. HPLC system diagram. Adapted from the literature18. ............................................................ 8

Figure 6. Taylor-Aris dispersion in laminar flow where (a) initial analyte plug and (b) the plug after time

t in the absence of diffusion; (c) is the plug after time t with a finite axial diffusion where the effect of

diffusion is minor as compared to the dispersive effect of the flow. (d) represents the plug after time t

with finite radial/transverse diffusion. Adapted from the literature34. ..................................................... 12

Figure 7. Fabrication process of: (1) aluminium hard masks; (2) SU-8 negative photoresist two level

mold; and (3) PDMS structures. The photoresists 1 and 2 should be selected to allow the spin-coating

of a thinner and thicker SU-8 layer, respectively. .................................................................................. 14

Figure 8. Structure of poly(dimethylsiloxane). ...................................................................................... 15

Figure 9. A 3D scale diagram of several pneumatically actuated valves in a peristaltic pump

configuration. The channels are 100 µm wide and 10 µm high. Adapted from the literature39. ............ 15

Figure 10. Chemical structure of ochratoxin A. .................................................................................... 19

Figure 11. Chemical structure of aflatoxin B1....................................................................................... 20

Figure 12. Sequence of steps involved in the fabrication of the aluminium hard masks. .................... 22

Figure 13. Sequence of steps involved in the fabrication of the SU-8 negative photoresist mold (iteration

1, 2 and valves layer molds) and of SU-8 negative and AZ 40 XT positive photoresist molds (iteration 3

and 4 fluidic layer molds). ...................................................................................................................... 24

xvi

Figure 14. Sequence of steps involved in the fabrication of the PDMS channels for the different

iterations, using distinct fabrication methods: soft lithography with irreversible sealing done by plasma

treatment (iteration 1 and 2); multilayer soft lithography and sealing using an adhesive PDMS layer

(iteration 3 and 4A) or plasma treatment (iteration 4B). ........................................................................ 25

Figure 15. Experimental setup for compressed air packing and valves actuation. (A) Switch buttons for

on/off actuation on the valves. (B) and (C) PCB for controlling of valves actuation and respective power

source. (D) Syringe pump. (E) Microfluidic device connected to the system by capillary tubing. ......... 29

Figure 16. Comparison between the first two CAD iterations created using AutoCAD. (1-A) corresponds

to a close-up representation of the bead trapping feature near the outlet where the channel has two

different heights: 100 and 20 µm. (1-B) corresponds to a microscopic image of the master mold

fabricated with the serpentine separation channel with a total length of 30 cm. (1-C) corresponds to the

microscopic image of the master mold where the bead trapping can be seen. (2-A, B) highlight the

simple cross-channel injector representation introduced into the design near the inlet and a microscopic

image of the injector on the master mold, with the two channels of different heights (100 and 20 µm).

(2-C) Microscopic image of the bead trapping feature and serpentine separation channel with a total

length of 12 cm. ..................................................................................................................................... 33

Figure 17. Comparison between the third and fourth CAD iterations created using AutoCAD. (3-A)

corresponds to a close-up representation of the cross-channel injector with integrated pneumatically

actuated valves. (3-B) corresponds to a microscopic image of the valves mold fabricated showing a

single valve channel. (3-C) corresponds to the microscopic image of the master mold where the

separation channel and cross-channel injector can be seen. (4-A, C) A close-up representation and

microscopy image, respectively, highlights the connection channel (height: 20 µm) designed between

the new cross-channel injector (height: 35 µm) and the separation channel (height: 100 µm) which

entraps beads within the separation channel. (4-B) Microscopic image of the new incorporated cross-

channel injector with integrated pneumatically actuated valves............................................................ 35

Figure 18. Comparison between the two injectors from iteration 3 and iteration 4. (A) Microscopic image

of the cross-channel injector and separation channel from iteration 3, packed with silica beads. (B)

Microscopic image of the cross-channel injector, connection channels and separation channel from

iteration 4, packed with silica beads. ..................................................................................................... 36

Figure 19. Microscopic images PDMS microfluidic structures (iteration 3) comprising valves of different

thicknesses originated by different spin-coating speeds, and its actuation. (A, B and C) are microscopic

images of valve channels spin-coated at 300, 400 and 500 rpm originating decreasing valves layer

thickness. (D and E) are microscopic images of the actuation of valve channels at P = 0 and 0.1 MPa,

respectively, for a 300 rpm spin-coated layer. (F) is a microscopic image pf the actuation of valve channel

at P = 0.1 MPa for a 400 rpm spin-coated layer. Green food colouring was flowed through the injector

channel for better visualization of valves actuation. .............................................................................. 37

Figure 20. Schematics illustrating the two strategies used for sealing of the microfluidics structures

comprising valves (iteration 3 and 4), namely bonding through half-cure and PDMS glue or bonding with

conventional plasma cleaner treatment. ................................................................................................ 38

xvii

Figure 21. Schematics comprising a timeframe of the beads trapping feature with packed silica beads

(left); a comparison between the different packing methods used throughout the experimental work

(centre); and a set of microscopic images of a fully packed 12 cm length separation channel with C18

bonded silica beads (right, A-D). ........................................................................................................... 41

Figure 22. Separation of a green coloured mixture. (A-C) corresponds to an illustration of the sequential

steps previously done to insertion of the mobile phase, including beads packing (A), green mixture

solution loading (B) and elution with Milli-Q water as the mobile phase (C). The microscopic images (i-

iv) show the progression of the separation throughout 10 minutes. (i) Initial sample plug starts to be

eluted with Milli-Q water as mobile phase. (ii) Methylene blue starts to be retained in the column. (iii)

Yellow plug travels through the column. (iv) Eluted components leave the column in a yellow plug with

traces of green. ...................................................................................................................................... 43

Figure 23. Schematics illustrating the two employed strategies for separation channel regeneration: (1)

increasing the ionic strength of the mobile phase or (2) protonation of the carboxymethyl group of the

stationary phase. ................................................................................................................................... 44

Figure 24. Plots comprising the fluorescence signal variance in function of the concentration of OTA

(top) and AFB1 (bottom) toxins. The correspondent microscopic images for the different concentrations

for each toxin are displayed on the left of the graphics. Fluorescence images were acquired at the

beginning of the microchannel with an exposure time of 1 second and a gain of 1x. ........................... 46

Figure 25. Elution profiles for toxins OTA and AFB1 previously adsorbed in a C18 bonded silica packed

microchannel. Elutions were performed during 10 minutes and fluorescence images acquired every 30

seconds at the beginning of the microchannel with an exposure time of 1 second and a gain of 1x. .. 47

Figure 26. Cross-channel injection and elution of the toxins AFB1 and OTA in reverse-phase separation

channel. (A) illustrates the mode of operation for loading and elution of the toxins using iteration 2 as a

microfluidic structure. (B) are fluorescence acquisitions in the beginning of the separation channel

comprising the cross-channel injector, of the AFB1 6 µg/mL elution with 100% CH3CN (v/v) during the

first 2 minutes. (C) are fluorescence acquisitions in the beginning of the separation channel, of the

elution with CH3CN:water (25:75 v/v) for the separation of OTA and AFB1, at a concentration of 6 µg/mL

Fluorescence images were acquired at the beginning of the microchannel with an exposure time of 1

second and a gain of 1x. ....................................................................................................................... 48

Figure 27. Schematics showing the operation of the cross-channel injector with integrated valves for

the injection and elution of the FITC plug by illustrating the sequential steps: washing (step 1), loading

(step 2), injection (step 3) and elution (step 4). Arrows indicate the direction of fluid flow and dark lines

the valves that are closed. ..................................................................................................................... 51

Figure 28. Sequential fluorescent image acquisitions of a FITC plug injection (i-iii) and progression (iv-

viii) throughout the separation channel without beads packed. Images were acquired with an exposure

time of 800 milliseconds and 3x gain. ................................................................................................... 52

Figure 29. Sequential fluorescent image acquisitions of a FITC plug injection (i-iii) and progression (iv-

viii) throughout the separation channel with beads packed. Images were acquired with an exposure time

of 800 milliseconds and 3x gain. ........................................................................................................... 53

xix

List of Acronyms

AFB1 - aflatoxin B1

OTA – ochratoxin A

FITC – fluorescein isothiocyanate

CE – capillary electrophoresis

μTAS – micro total analysis system

LOC – lab-on-chip

LC – liquid chromatography

HPLC – high performance liquid chromatography

GSC – gas solid chromatography

GC – gas chromatography

SFC – supercritical fluid chromatography

LLC – liquid-liquid chromatography

MEKC – micellar electrokinetic chromatography

LSC – liquid solid chromatography

SEC – size exclusion chromatography

IEC – ion exchange chromatography

AC – affinity chromatography

BPC – bonded phase chromatography

RPC - reversed-phase chromatography

NPC - normal-phase chromatography

NARP - non-aqueous reversed phase chromatography

HILIC - hydrophilic interaction chromatography

IPC - Ion-pair chromatography

MEMS - microelectromechanical systems

PDMS – polydimethysiloxane

DWL - direct laser writer

MSL - multilayer soft lithography

IPA – isopropanol

DI – deionized

PGMEA - propylene glycol ether acetate

CAD - computer assisted design

PEG - polyethylene glycol

PCB - printed circuit board

1

1. Introduction

1.1. Motivation

Separation of mixtures is very important in synthesis, industrial chemistry, biomedical field and

chemical analyses. A separation process can be defined as the method to achieve any phenomenon

that converts a mixture of chemical substances into two or more distinct product mixtures, at least one

enriched in one or more of the mixture’s constituents. It is also normally associated with the removal of

impurities to achieve purification of the product, depending on the purpose: either preparative in the

case of purification - as just exemplified - or analytical. The goals of an analytical separation are usually

to eliminate or reduce interferences so that quantitative analytical information can be obtained from

complex mixtures. Many separation methods exist nowadays, ranging from precipitation1 and extraction2

to various chromatographic techniques3,4. All of them differ in their separation principle that can be based

on chemical or physical properties such as size, shape, mass, density or chemical affinity between the

components of the mixture. Apart from the separation itself, either analytical or preparative separations

require a detection mode and later translation of signal into real information. To note that in a technique

such as chromatography, quantitative information is obtained nearly simultaneously with the separation

unlike other procedures already mentioned above.

Analytical separations can occur at any scale: from a laboratory scale to the industrial scale.

However, there is an increasing demand for smaller and smaller instrumentation for analytical

separations5, requiring high-throughput answers to its problems but more importantly, fast and on-spot

analyses of products of interest. This implies small sample and set-up size, low costs and easy disposal

of wastes.

Microfluidic devices have been the answer for these demands in the last years. Although some

work had already been reported6, it was since capillary electrophoresis (CE) was demonstrated in

microfabricated channels in the year of 1992 for the first time7, that a spurt of interest in microfluidic

devices has been generated in analytical separations field. The reason miniaturized systems are gaining

popularity is obvious: size matters. A miniaturized system can attain a high compactness of components

and this simplifies multiplexed systems for providing high sample throughput. Single analysis can as

well be considerably accelerated with the reduced setup sizes because distances for molecules to travel

are drastically shortened – sample size is therefore minimized. The same applies for costs of the device

as less sample is required. More importantly, all the steps involved in the process – from sample

preparation to signal acquisition – can be integrated onto a single device. This concept was first

described in 1990 as micro total analysis system (μTAS)8, nowadays more known as Lab-on-Chip (LOC)

devices9.

2

One of the most popular separation methods – liquid chromatography (LC) – has been one of the

most prolific applications in LOC devices. However, efforts directed on developing pressure driven

separations (analogous to conventional liquid chromatography columns and HPLC – High Performance

Liquid Chromatography) in a chip based platform have been pursued by a minor number of research

groups10,11,12. This can largely be explained by the much different technical challenges involved in

successful execution, such as on-chip integration of the porous stationary phase, ultra-low volume

sample injection, adequate chip sealing, on-chip pumping, and on-chip detection. Nevertheless, non-

stop developments in microfabrication and nanotechnology, as well as propagation of wafer level

processing well outside the microelectronics industry, created an abundant ground for exploration of

such possibilities with high chance of future innovations. Given the current circumstances of the field

stated above, it is imperative to take advantage of the new technologies available to create a

chromatographic chip which not only meets the goal of fully integration like attempted previously12, but

also aims for a simpler fabrication process (and therefore more economically viable), user-friendly

operation and versatility for a broad range of analytes/applications.

Within the range of relevant analytes for this device, mycotoxins are a particularly important

application since they contaminate the diet of a large proportion of the world’s population, especially in

low-income countries where the contamination levels are the highest13. They represent a serious global

health issue as per most known mycotoxins, like OTA and AFB1, are potential carcinogenic (Group 2B)

and confirmed carcinogenic (Group 1), respectively, for humans14. The fact that such a device can have

a broad range of applications, including critical areas such as food safety, is of upmost importance for

the motivation of this work.

1.2. Chromatography

Chromatography is a broadly used method for the separation, identification, and determination of

chemical components in complex mixtures. No other separation method is as powerful and largely

applicable as chromatography. It is such a diverse method that it can be applied to several systems and

techniques3,4,15. However, all of them have common concepts and features associated that define what

is a chromatographic method. Therefore, a brief guide to the most important concepts will be addressed

in the following sections and represent an summary of the literature16.

1.2.1. General concepts

Chromatography is a technique in which the components of a mixture are separated based on

differences in the rates at which they are carried through a fixed stationary phase by a gaseous or liquid

mobile phase. The stationary phase is the phase that is fixed either in a column or on a planar surface.

On the contrary, the mobile phase is the one that moves through the stationary phase carrying with it

the analyte mixture. The mobile phase might be a gas, a liquid, or a supercritical fluid.

3

1.2.1.1. Elution

Elution is a procedure in which solutes are washed through a stationary phase by the movement

of a mobile phase. The mobile phase that departures the column is labelled the eluate.

16,17

In Figure 1, an example of a protein mixture being eluted by column chromatography is shown.

The column is packed with a matrix (solid, porous material) consisting on the stationary phase. Initially,

the mobile phase (eluent) is introduced onto the column, occupying the empty spaces existent in the

matrix. An eluent is a solvent used to transport the components of a mixture through a stationary phase.

Then, the solution containing the protein mixture is inserted as a narrow plug, corresponding to the time

t0 in the diagram. Next, elution happens by compelling the mixture through the column and continuously

adding new mobile phase. Given that the solute (in this case, the proteins) only moves with the help of

the elution, it is usually said that the average rate at which a solute migrates depends on the fraction of

time it spends in that phase, being small for solutes that are strongly retained by the stationary phase

(example: solute B in Figure 1). As shown in the previous figure, these different rates of migration will

result in individual bands along the column. The separation of each component involves

collecting/detecting these different bands as can be seen in the diagram above.

Figure 1. Column chromatography. Left - the standard elements of a chromatographic column include a matrix

supported inside a column, generally made of plastic or glass. A protein solution to be separated can percolate the

column (starts on top). Proteins migrate through the column at different rates because of different interactions

established with the matrix. Right - diagram representing the separation of the chromatography procedure illustrated

on the left. The detected signal at the various stages of elution is shown. Adapted from the literature16,17.

4

1.2.1.2. Chromatograms

The separated components can be collected at the end of the column and/or detected. But how

does this happen? By simply placing a detector that responds to solute concentration at the end of the

column (or other support, depending on the chromatography) when elution occurs, a series of peaks

can be obtained, if the signal is plotted as a function of time or added eluent volume. This plot in

chromatography is usually called chromatogram (Figure 2) and serves as both qualitative and

quantitative analysis.

16,17

The identification of each component on the mixture can be done by analysing the position of the

peaks maxima on the time axis. Also, the peak areas provide quantitative information of the amount of

each species present in the mixture. However, several times the chromatogram does not turn out so

well, due to band broadening, as seen in Figure 2 (right illustration). There are many possible methods

to overcome this difficulty that will be addressed further on: some of them consist in improving the band

separation rate or decrease the rate of band spreading by manipulating several variables that influence

solute migration rates.

1.2.1.3. Retention time

𝑡𝑅 = 𝑡𝑆 + 𝑡0 (Equation 1)

The retention time, 𝑡𝑅, (Equation 1) is the time between injection of a sample and the presence of

a solute peak at the detector of a chromatographic column: different solutes have different retention

times. In Figure 3, a chromatogram of a two-component (I and II) mixture is shown. As seen below, t0,

the dead time, is the time it takes for an unretained species to pass through a chromatographic column.

Figure 2. Chromatogram. Left - elution of components A, B and C is represented by plotting the detector signal

intensity in millivolts versus time of elution in minutes. Right - two-component chromatogram exemplifying two

approaches for improving separation: (a) original with overlapping peaks, (b) improvement by increase in band

separation, (c) improvement by decrease in band widths. Adapted from the literature16,17.

5

All components spend at minimum this total of time in the mobile phase. Separations are based on the

different times 𝑡𝑆, that components spend in the stationary phase.

1.2.2. Column performance

The most important factor when designing or assessing a chromatographic column is its

performance. More important than simply separating the components of a mixture, it is desirable that

this separation is efficient. To obtain optimal separations, sharp, symmetrical chromatographic peaks

must be obtained. This means that band broadening must be limited. However, there are many factors

imposing barriers to a highly efficient separation in chromatography. These factors will be addressed in

the following sections by briefly highlighting the models behind column efficiency assessment. More

information about this can be found on the literature18, since a more profound mathematical approach

to the models is not within the scope of this work.

1.2.2.1. The theoretical plate model of chromatography

There are two essential parameters in this model approach used to measure chromatographic

column efficiency: plate height, 𝐻, and number of theoretical plates, N. They are correlated by the

following equation:

𝑁 = 𝐿

𝐻 (Equation 2)

Where 𝐿 is the length of the column packing. As it can be deduced, the smaller the value for 𝐻

the higher the value for 𝑁, meaning a higher number of plate count, therefore achieving more column

efficiency. The plate model assumes that the column comprises many distinct layers, termed theoretical

Figure 3. Chromatogram of a two-component (I and II) mixture. t0 represents the dead time, tR1 and tR2 the

retention times of the components I and II, respectively. W1 and W2 are the band widths of each component and

Δt is the difference in elution time between the two components. Adapted from Thermopedia.

6

plates. Separate equilibrations of the sample between the stationary and mobile phase occur in these

"plates". The analyte moves down the column by transfer of the mobile phase from one plate to the next.

1.2.2.2. The rate theory and the van Deemter equation

Theoretical studies of zone broadening in the years around 195019 led to the van Deemter

equation (Equation 3). A more correct explanation of what occurs inside a column considers the time

taken for the solute to equilibrate among the stationary and mobile phase (contrasting the plate model,

which accepts that equilibration is infinitely fast). This means that the band shape depends on the rate

of elution. It also may be affected by the diverse paths accessible to solute molecules as they travel

between the stationary phase. This all translates into Equation 3, where 𝐶𝑆 and 𝐶𝑀 are mass-transfer

coefficients for the stationary and mobile phases, respectively, and 𝑢 is the linear velocity of the mobile

phase.

𝐻 = 𝐶𝑀𝑢 (𝑜𝑟 𝐴) + 𝐵

𝑢+ 𝐶𝑆𝑢 (Equation 3)

The term (𝐵𝑢⁄ ) represents longitudinal diffusion. This factor causes band width to rise with time,

and it happens whether or not the mobile phase is flowing. The time spent by the band during its way

through the column differs inversely with the flow rate, so the influence of longitudinal diffusion on band

width decreases for increased flows. In liquid chromatography, diffusion rates are much smaller in value

so this term has an almost insignificant influence on band broadening. On the other hand, the term (𝐶𝑆𝑢)

represents the stationary phase mass-transfer. Among the analyte molecules, there will be ones that

enter further into a particle pore than others, spending in it a given and varied time. Molecules that spend

less time in the particle will move further along the column, with a resulting increase in band width. This

contribution to band broadening increases as the flow rate increases. Also, when the band leaves the

column has to pass through the detector, resulting in additional peak broadening. Finally, the term A or

(𝐶𝑀𝑢) represents the mobile phase mass-transfer. Zone broadening in the mobile phase is caused by

the multiple paths a molecule can take through a packed column, especially when the lengths of these

paths can differ significantly. This phenomenon is usually called eddy diffusion. At low mobile phase

velocities, the molecules are not significantly dispersed due to the contribution of regular diffusion effects

that attenuates the eddy diffusion. At high velocities, there is no sufficient time for diffusion to act, and

then band broadening due to the different path lengths is seen.

1.2.3. HPLC

There are diverse kinds of chromatographic methods and these are classified according to the

shape of bed (planar or column), physical state of the mobile phase and separation mechanisms.

Furthermore, there are other adapted arrangements of these chromatographic techniques relating

7

different mechanisms and are therefore considered as modified or specific chromatographic

techniques20. In the following diagram (Figure 4), the techniques are organized by the mobile phase

physical state, stationary phase and mechanism of separation.

As seen in Figure 4, LC is the one with broader range of available separation mechanisms. This

versatility and the rapid expansion of HPLC since the early 1970s21 has put liquid chromatography on

top of the most important analytical separation techniques.

Early liquid chromatography used to be performed in columns with an internal diameter from 10

to 50 mm and were around 50-500 cm long, therefore particle size used to be around 150-200 µm to

ensure a reasonable flow rate was achieved without a high demand on pressure. Together with the early

development of the theory of liquid chromatography, it was understood that in order to decrease the

plate height smaller particle size (3-10 µm) was in need21. Of course, this new feature required higher

pumping pressures along with the development of new detectors for continuous monitoring. Throughout

that time, each procedure presented key enhancements in convenience, speed, resolving power

detection, quantification and applicability to new sample natures. It is easy to see how HPLC embodies

the modern culmination of the progress of liquid chromatography. Consequently, the equipment for

HPLC nowadays tends to be considerably more elaborate and therefore expensive, time-consuming

and requiring specialized personnel for operation.

1.2.3.1. Instrumentation

Within the scope of this work, it is important have in mind the main features of HPLC

instrumentation to understand the need of several components integrated in the microfluidic device and

even assess the superiority of such device when compared to HPLC.

Figure 4. Diagram of the chromatographic methods. GSC – gas solid chromatography, GC – gas chromatography,

SFC – supercritical fluid chromatography, LC – liquid chromatography, LLC – liquid-liquid chromatography, MEKC

– micellar electrokinetic chromatography, LSC – liquid solid chromatography, SEC – size exclusion

chromatography, IEC – ion exchange chromatography, AC – affinity chromatography and BPC – bonded phase

chromatography.

8

18

In Figure 5 a HPLC system diagram is presented where the main components are highlighted,

such as the solvent reservoir followed by an high-pressure pump and a autosampler (injector) just before

the chromatographic column. Sometimes, it is important to maintain a specific temperature for enhanced

column performance, therefore we can observe a column oven placed on the HPLC system diagram.

Following the column, there is a detector connected to a system that will analyse the signal and

transform it into a chromatogram.

More specifically, the mobile phase reservoirs are often accompanied with components placed

before the pump to remove dissolved gases (in-line degassing) which normally lead to irreproducible

flow rates and band spreading. Primarily, it is important to improve the pump operation to ensure fixed

flow rates, but dissolved oxygen can also be a source of background noise for the detector22. Also,

provisions should be included to remove dust and bubbles that usually interfere with the signal

acquisition of most detectors, especially when HPLC-grade solvents are not used. Regarding tubing

included on the HPLC equipment, care should be taken to ensure that tubing length and diameter are

chosen to minimize peak-broadening contributions, most precisely in tubing where the sample is

present: between the autosampler and the column, and between the column and detector23.

There are several requirements for most HPLC conventional pumping systems: they must

generate high pressures up to 6000 psi, have a pulse-free output and have a good flow reproducibility.

Most commercial instruments use the reciprocating pump type which meets the requirements stated

above: capacity for high output pressure (up to 10000 psi), achieves constant flow rates independent of

column back-pressure and solvent viscosity, in addition to small internal volumes and good adaptability

to gradient elution. Finally, most of these pumps are computer-controlled devices to measure the flow

rate and rapidly adapting the settings to maintain a constant flow rate18.

Figure 5. HPLC system diagram. Adapted from the literature18.

9

Before injection of the sample and mobile phase into the column, it is important to consider the

type of elution needed for the given separation. In an isocratic elution, the solvent composition remains

constant throughout the separation and can either be a single solvent or a mixture. However, gradient

elution is known for improving the separation efficiency and consists in changing the composition of the

solvent throughout the separation either continuously or in a step-wise fashion. Although for isocratic

elution the mobile phase can be hand-mixed, most of the times the HPLC system can have on-line

mixing integrated in the system both for isocratic elution and gradient elution, where on-line mixing is

required. In many HPLC systems, proportioning valves are used for this purpose. Sample injection onto

the column is a very important aspect since sample size can impact peak broadening as well the need

for an automated injection system (autosampler) to increase the throughput of analysis. A feature widely

also widely use is a sample loop that provides a choice of sample sizes ranging from 1 to 100 µL or

more18,23.

Most analytical columns used in HPLC nowadays have a length from 5 to 25 cm, being the most

common lengths 10-15 cm long with an internal diameter of 4.6 mm and packed with 5 µm beads. Apart

from the analytical column, HPLC systems usually have precolumns: placed between the mobile phase

reservoir and injector, a scavenger column minimizes losses of stationary phase from the analytical

column by ensuring the mobile phase is already saturated with silicic acid from the silica packing. In

addition, another precolumn commonly used is a guard column – placed between the injector and the

column - that has a similar packing to the analytical column and therefore prevents impurities to

contaminate the analytical column and increases its lifetime. Finally, post-column we have a detector

that ideally would be small, compatible with liquid flow and universal. The restriction in size advents for

the need for low internal volume in order to minimize extra-column band broadening. Unfortunately,

there is no detector that meets all the requirements and is at the same time highly sensitive, mostly

because the detector used will depend on the nature of the sample under analysis18.

1.2.3.2. Chromatographic modes

In Table 1, several types of operation modes on HPLC are shown. One of the most relevant is

the reversed-phase chromatography (RPC) and its opposite, normal-phase chromatography (NPC).

They are distinguishable based on the relative polarities of the mobile and stationary phases.

In the beginning, most separations were based on highly polar stationary phases (such as silica)

and a relatively nonpolar solvent (hexane) was the mobile phase. This type of chromatography is now

called normal-phase chromatography. The less polar molecule is eluted first and increasing the polarity

of the mobile phase will decrease the elution time. On the other hand, in reversed-phase

chromatography, the stationary phase is nonpolar, often a hydrocarbon brushlike surface, and the

mobile phase is a relatively polar solvent (such as water, methanol, acetonitrile). The most polar

component elutes first, and decreasing the mobile phase polarity will decrease the elution time16.

Nowadays, the majority of HPLC separations are performed in this chromatographic mode due to both

10

the variety of stationary phases available and the broad range of molecules that can benefit from the

separation4,24.

Table 1. HPLC chromatographic modes are summarized with attached examples present in the literature.

Mode Description Literature example

Reversed-phase chromatography (RPC)

The column is nonpolar and the mobile phase is a polar mixture of water plus organic solvent; RPC is the most widely used mode, especially for water-soluble samples.

Bonfatti et al.4

Normal-phase chromatography (NPC)

The column is polar and the mobile phase is a mixture of less-polar organic solvents; NPC is used mainly for water-insoluble samples, preparative HPLC, and the separation of isomers.

Panfili et al.25

Non-aqueous reversed phase chromatography

(NARP)

The column is nonpolar, and the mobile phase is a mixture of organic solvents; NARPC is used for very hydrophobic, water-insoluble samples.

Lísa et al.26

Hydrophilic interaction chromatography (HILIC)

The column is polar, and the mobile phase is a mixture of water plus organic; HILIC is useful for samples that are highly polar and therefore poorly retained in RPC.

Bajad et al.27

Ion-exchange chromatography (IEC)

The column contains charged groups that can bind sample ions of opposite charge, and the mobile phase is usually an aqueous solution of a salt plus buffer; IEC is useful for separating ionizable samples such as acids or bases, and especially for the separation of large biomolecules.

Rea et al.28

Ion-pair chromatography (IPC)

RPC conditions are used, except that an ion-pair reagent is added to the mobile phase for interaction with sample ions of opposite charge; IPC is useful for the separation of acids or bases that are weakly retained in RPC.

Ibáñez et al.29

Size-exclusion chromatography (SEC)

An inert column is used with either an aqueous or organic mobile phase; SEC provides separation based on molecular weight and is used mainly for large biomolecules or synthetic polymers.

Gellein et al.30

Chiral chromatography

A chiral agent is fixed on the surface of a solid support and then modes of interaction can happen, as for example, based on attractive forces between each other or the fitting into chiral cavities, forming inclusion complexes.

Peng et al.31

Another relevant chromatographic mode for this work and present in Table 1 is the ion-

exchange chromatography. The separation principle is based on molecule’s charged groups: the column

retains molecules based on the electrostatic interactions with opposite charges from functional groups

present in the stationary phase. Analyte molecules will have to compete with counterions present in

solution for binding to the functional groups so that initially, molecules that do not bind or bind weakly to

the stationary phase are washed away. Elution of bonded molecules will happen when the mobile phase

is altered to provide an increase in the concentration of the counterions that will compete with the bonded

molecules. This alteration in counterions concentration can be done by changing either the ionic strength

or the pH of the mobile phase. The stationary phase can be classified in two types: when it features

positive groups, it is called an anion exchanger and when there are negative groups it is a cation

exchanger. Finally, it can also be classified as strong or weak exchanger based on whether it can

maintain its charge within a wide range of pH (strong) or only maintaining the charge in small pH

interval15.

The other listed HPLC chromatographic modes also play an important role in the widespread

application of HPLC in chemical analysis, but will not be the focus of this work.

11

1.3. Microfluidics

MEMS (microelectromechanical systems) is a field that was born after the need for

miniaturization of all kinds of systems surged, allowing the creation of devices that are normally between

the range of 1-300 µm in size. Fluid flow in these conditions revealed phenomena worth exploring a new

field that we today know as microfluidics which can be defined as “(…) the study of flows that are simple

or complex, mono- or multiphasic, which are circulating in artificial microsystems (…)”, according to the

literature32. Since technologies for MEMS were established (mostly silicon-based devices) - around the

1980s – there was already enough knowledge to start working on microfluidic devices. However, it was

only after 1992 that microfluidics was recognized for numerous potential applications, particularly in

chromatography and CE7. Nowadays, scientists are very interested in microfluidics, given that it is a way

simpler than MEMS, by not almost never including moving components and being “soft-based”, as

polydimethysiloxane (PDMS) is the main component of nearly all devices32. In this section several

aspects of microfluidics will be addressed, such as general concepts of physics at the microscale and

fabrication as well as the implementation of chromatographic separations at the microscale.

1.3.1. General concepts

To understand the physical phenomena prevailing in miniaturized systems the scaling laws must

be studied. A scaling law can be defined as how a certain physical quantity varies with the size of the

system under analysis. Analysing Equation 4, the concept can be understood:

𝑈 ~ 𝑏2∆𝑃

𝜇𝐿 (Equation 4)

Where 𝜇 is the fluid viscosity, Δ𝑃 the pressure difference, 𝐿 the length of the channel and 𝑏 the

transverse dimension. Flow velocity, 𝑈, is equal to l1. Because there are so many variables into

consideration, some assumptions must be made in order to predict the scaling law applicable. In this

case, pressure difference of a microfluidic channel was considered to be constant. Everything else must

be arranged considering the microfluidic context, as for example the laminar flow regime that is

characteristic of microfluidic operation. Generally, the physical quantity that has a lower exponent when

compared to others is the one that will become dominant at the microscale (for example, capillary forces

winning over gravitational forces). Regarding microhydrodynamics, it is also important to highlight that

generally, liquid flow in microchannels is controlled by the dimensionless Reynolds number (Equation

5):

𝑅𝑒 = 𝜌𝑈𝐿

𝜇 (Equation 5)

12

Where 𝑈 is the velocity of the fluid, 𝜌 the density, 𝐿 the characteristic linear dimension and 𝜇 the

viscosity of the fluid. Considering that fluid velocities are very small and that dimensions of channels are

around the micrometre range, the values for Reynolds number are usually between 1 to 10 units which

falls under the laminar flow regime (Re < 2000).

As already mentioned, the need for smaller particle sizes in chromatography led to higher

operating pressures, creating the concept of HPLC. Such correlation can be addressed by Equation 6:

∆𝑃 ~ 𝐿2𝜇

𝑏2𝑡𝑅 (Equation 6)

Where L is the length of the column and b the typical size of the space between grains or pores.

𝑡𝑅 is the retention time of a given analyte. If we assume that all the variables are constant except for b

(particle size), one can observe that ΔP will increase substantially as b decreases.

Finally, when dealing with pressure-driven flows within a microchannel, an interesting effect is

the one of Taylor dispersion, where a shear flow can increase the effective diffusivity of a given molecule.

Shear acts disperse the concentration distribution in the direction of the flow, increasing the rate at which

it spreads. The overall effect is an enhancement of axial dispersion over molecular diffusion alone in the

absence of flow. There is also a significant difference in the spreading behaviour of the solute shortly

after its injection and after several diffusion times have elapsed after its injection33. In Figure 6, the

spreading behaviour shortly after its injection time can be observed. The cross-channel diffusion is yet

to happen and the distance between the fastest moving molecules at the center and the slow particles

on the wall stretches linearly with time. After some time, enough to let cross-channel diffusion to act, the

edges of the paraboloid change to form a blob which is nearly uniform across the channel and the plug

spreads.

34

Figure 6. Taylor-Aris dispersion in laminar flow where (a) initial analyte plug and (b) the plug after time t in the

absence of diffusion; (c) is the plug after time t with a finite axial diffusion where the effect of diffusion is minor as

compared to the dispersive effect of the flow. (d) represents the plug after time t with finite radial/transverse diffusion.

Adapted from the literature34.

.

13

However, chromatographic processes, such as mass exchange between phases or nonuniform

fluid flow – which were previously described in the van Deemter equation - can greatly affect the

effectiveness of molecular diffusion in causing dispersion, so each microfluidic separation channel has

to be regarded individually and not much generalizations can be done in assessing the role of this effect

on separation efficiency. Additionally, it is not only the effects of pressure-driven flow, packed bed and

chosen chromatographic conditions that can influence plug spreading along the channel, but also factors

such as channel geometry, with reported data on turns35,36 and corners37 and different cross-sections

geometries33 influencing plug spreading.

1.3.2. Microfabrication

In this section, some of the most common microfabrication techniques currently used for

microfluidic operation will be addressed. Special focus will be given to the technologies related to the

work in progress, namely photolithography and soft-lithography technologies.

1.3.2.1. Photolithography

Photolithography is a procedure used in microfabrication to pattern on a substrate. By the action

of light, it transfers a pattern to a photosensitive chemical, usually called a photoresist, previously coated

on the substrate. This pattern can be defined by direct laser writer (DWL) system or by a previously

patterned mask. Optical lithography is the most used technology and comprises wavelengths between

300 and 450 nm32. Masks are generally made of glass substrate on which aluminium (or a metal with

similar purpose) is usually deposited along with a photoresist to be further patterned using a DWL

system. Finally, it is required to etch by wet etching the aluminium that is not protected by the photoresist

to obtain the final features. The mask should be the object with the most precision possible, as the

following substrates will never have more precision than the mask itself. Masks are very useful to create

reusable molds that contain the same features as the masks and are further used in soft-lithography

fabrication.

To fabricate a mold, a photoresist is spin-coated on a silicon substrate. While the resist spreads

on the substrate, the solvent does not evaporate totally, making the film look like a soft solid. To remove

the remaining solvent, the polymer is heated before the exposure step. For exposure, the photoresist

coated substrate is aligned with the mask and the incident light starts chemical reactions in the

photoresist, which will alter the solubility in certain solvents. For example, for a positive resist the zones

exposed to the light become soluble and disappear while for a negative resist the zones exposed to the

light become insoluble in solvent, therefore staying in the substrate. It is important that the photoresists

used have a great disparity between the solubility constants of the exposed and unexposed portions,

14

high photosensitivity and high resistance to some chemicals used in afterward steps of

microfabrication32. The full fabrication of mold (including mask fabrication) is highlighted in Figure 7:

1.3.2.2. Soft lithography technologies

PDMS plays a very important role in microfluidics as one of the elastomeric materials in

popular use in soft lithography due to great capacity for rapid prototyping and easy fabrication without

expensive equipment. PDMS belongs to a family of polymers that especially contain silicon oils as seen

in Figure 8. When PDMS reaches a temperature higher than the polymerization temperature (around

70 ºC) and is mixed with a reticulating agent, it forms an elastomer whose main properties are: optical

transparency (the UV spectrum permits the visualization of flows), elasticity (allows the fabrication of

valves and pumps), good insulation, low surface energy (easier to peel off the PDMS form the mold),

chemical inertia, non-toxicity and low permeability to water (but permeable to gas)38. Also, untreated

PDMS is hydrophobic, and becomes temporarily hydrophilic after being submitted to oxidation of the

surface by oxygen: plasma creates silanes by oxidation of methyl groups. This oxidised PDMS adheres

to glass, silicon or even PDMS itself, if the surfaces receive the same treatment38. To achieve structures

like microchannels a technique called replica molding is used: a mold made of a hard material is

necessary (normally made of silicon or deposited photoresist) and a mixture of PDMS and reticulating

agent is poured on the mold. The mixture is baked and the PDMS polymerizes/reticulates, becoming

Figure 7. Fabrication process of: (1) aluminium hard masks; (2) SU-8 negative photoresist two level mold; and

(3) PDMS structures. The photoresists 1 and 2 should be selected to allow the spin-coating of a thinner and

thicker SU-8 layer, respectively.

15

solid. The last step consists on peeling off the PDMS structure from the mold and sealing against glass

or a PDMS slab to allow liquid flow in the channels.

After molding, the precision of PDMS structures is quite good – around submicrometric

values – but if the elastomeric character of the material and aging phenomena is taken into account, the

best channel dimensions are usually between 5 and 500 µm. Indeed, it is possible to fabricate structures

with higher aspect ratios but a risk is taken in whether or not these structures will collapse due to the

deformability of PDMS32.

Besides the advantages already mentioned, PDMS also allows the fabrication of integrated

pumps and valves by a technique developed by Unger et al designated multilayer soft lithography

(MSL)39,40. This technique combines soft lithography with the capability to bond multiple patterned layers

of elastomer and sealing them through half-curing of each patterned layer and baking them again

stacked together to complete the curing process. The principle is to have two opposite (the ratio of

element A and B in the elastomer mixture is substantially unalike) layers in contact with each other since

reactive molecules remain at the interface between the layers and can be further cured to produce a

sealing. More layers can be added by simply repeating this procedure. The valves were fabricated using

a cross-channel architecture, as seen in Figure 9, and had an actuation area of 100 x 100 µm in thin a

layer with a thickness of about 30 µm, designated control layer. When pressure was applied to the

control channel filled with water, this membrane would deflect to close the lower fluidic channel

beneath39. This new technology allowed for the incorporation of valves within PDMS microchips with

almost zero dead volume, a very interesting aspect in many applications, as for example the integration

of chromatographic separations in microfluidic devices. More recently, many iterations of these valves

or even new type of features have been integrated into chips using multilayer soft lithography, from

magnetic actuated valves41 to pneumatic valves in folded 2D and 3D fluidic devices made from plastic

films and tapes42.

39

Figure 8. Structure of poly(dimethylsiloxane).

Figure 9. A 3D scale diagram of several pneumatically actuated valves in a peristaltic pump configuration. The

channels are 100 µm wide and 10 µm high. Adapted from the literature39.

16

1.3.3. Chromatography-on-chip

Chromatography on chip is not something new but not an entirely finished business either. Since

CE was demonstrated in microfabricated channels in 1992 for the first time, a burst of interest in

microfluidic devices has been generated in analytical separations field7. LC has been one of the most

prolific applications in LOC devices, along with CE. However, efforts directed on developing pressure

driven separations (analogous to conventional liquid chromatography columns and HPLC) in a chip

based platform have been pursued by a minor number of research groups, even less when referred to

packed columns11,12,43,44,45 in the past two decades. This is mainly due to the technical challenges

involved in successful execution, such as on-chip integration of the porous stationary phase, on-chip

ultra-low volume sample injection and pumping and on-chip detection which will be addressed in this

section as well as the recent advances that are now enabling the chromatographic integration on

microfluidic devices to rise again.

1.3.3.1 Column design and fabrication considerations

In a HPLC-like microfluidic device, column and particles sizes must be scaled down at the same

time. Yet, to accomplish speed and separation efficiency that equals the ones from conventional HPLCs

the chip has to be submitted to high pressures, not only for the small particles and channel dimensions

used, but also for the need of increased channel length, which will cause more hydraulic resistance.

This demand causes enormous technical complications in execution of HPLC-like microchips that can

support such high pressures without leaking. Alternatively, if pressure stays identical, the separation

efficiency is worsened in the same degree and no enhancement in speed is gained, when related to

conventional HPLC. This is only one of the reasons why since the expansion of microfluidics in

chromatography, mainly CE has been integrated in microchips. Most importantly, the big advantage of

pressure-driven flow is that it is largely independent of the chemistry of the separation principle46.

To achieve separation efficiency, the channel size, shape and in particular, the aspect ratio must

be carefully designed not only considering the pressure requirements. For example, in packed channels,

the ratio of channel width/depth to particle size must be considered to avoid the known “wall effect”

where quality/density of packing or flow behaviour will be severely different from the bulk channel46,47.

This effect is more pronounced in microfluidic devices where the ratio is normally around 10. For values

much higher than 10 the wall region will only constitute a small part of the channel, and therefore the

impact of this effect will be diminished. On the other hand, for values much lower than 10 the wall and

the bulk region become very similar, therefore such effect loses its relevance. Different cross-sectional

shapes can also affect the separation efficiency by causing dispersion, either by impacting the packing

density and order or the fluid flow as corner regions tend to be of stagnant47.

17

Relatively to stationary phase considerations, there are three main ways to insert a stationary

phase into a channel: covering the walls of the channel with active elements of the stationary phase

(open channel chromatography)48; prepare packed beds of particles that will retain the analytes and

finally, creating monoliths which either already include the retentive elements or can be fixed with it later.

Open channel chromatography is the simplest way to perform the integration of chromatography into

chips, the big advantage is the ease of preparation of the column and also the low pressure required to

flow the mobile phase through the channel, even though small depths are required to compensate the

low diffusion coefficients and mass transfer to occur. However, a big disadvantage is the low surface to

volume ratio, which will decrease the capacity of processing a reasonable volume of sample and

therefore only the most sensitive methods of detection can be coupled to the device49.

There is also the possibility of simply packing the channel with porous media as usually done in

conventional LC. The big benefit of using packed channels is that essentially all the particles that have

been developed over the years for LC can be employed. The particle size should never be the only

factor in consideration, some other important aspects when designing the packed bed are: size

dispersity, porosity of the particles, and additionally, the pressure capacity of the microchip. Considering

all these factors, it seems that the degree of order in the packing is the biggest challenge on the way to

perfectly packed beds and the answer lied within the packing technique49. Also, some structure/material

is necessary to keep the beads trapped inside the channel, both during packing and operation: many of

the solutions used until now can cause band broadening and, consequently, reduce the performance of

the separation. Some examples of currently used designs are a cage-like PDMS device50, renewable

and pneumatically controlled pillar array51 or a double weir design52.

Instead of pressuring particles into a channel, a different alternative is to synthetize a porous

monolithic structure inside the channel. The main advantage is to avoid high packing pressures as well

as tricks to keep the stationary phase within the channel. However, the batch-to-batch reproducibility of

these monoliths is very hard to achieve within the acceptable range, not to mention the monolith

fabrication technology is still way too complex and undiscovered to be reasonably dominated soon and

widely implemented on chips49.

1.3.3.2 Injection and sample volume considerations

Another consideration to take into account is whether or not valves can be integrated on chip,

which will for sure impact on how “portable” can such a device be. Many approaches have been

proposed both for valves incorporation to optimize and integrate the plug injection into the

chromatographic column. One of the first components to tackle is the injector: there are many types of

designs for injectors that can be used nowadays. The most simple ones are the “double-T” junction53 or

a cross-channel design43 at the column inlet. They are preferable since they do not require valves and

can generate small sample plugs. The main disadvantages of these injectors are their propensity to

display sample dilution and leaks that can cause broadening of the plug previously to the insertion on

18

the channel due to backpressure issues. Considering the difficulties, more sophisticated designs of on-

chip injectors are available12 which often incorporate valves within the device or bulky instrumentation

outside the device. Ideally, the incorporation of valves would be preferable since it would offer zero dead

volume injections.

1.3.3.3 Literature examples on chromatography-on-chip

Some examples of different approaches to the chromatography-on-chip approach present in the

literature, are referred in this section. The first example is a highly efficient and ultra-small volume

separation by pressure-driven liquid chromatography in extended nanochannels10. In this work, the first

pressure-driven liquid chromatography system that allows separation of atto- to femtoliter sample

volumes was designed. Furthermore, the separation efficiency was very high and only within a few

seconds. In comparison to a conventional packed HPLC column, the separation is much faster (by 2

orders of magnitude), has a smaller injection volume (9 orders), and a higher separation efficiency by 1

order of magnitude. However, this microchip uses an open-column which means flexibility in relation to

the stationary phase is lost and also the fabrication process is more complicated than soft lithography

processes.

The next example is the work reporting the fabrication of high-quality microfluidic solid-phase

chromatography columns44. In this work, the authors present a column geometry that allows strong and

high-yield packing of chromatographic channels fabricated using MSL technique that can be

incorporated in an integrated microfluidic system for HPLC, providing several prospects for analytical

and preparative applications where low sample consumption, low cost, and automation are important.

However, this packing method adds complexity to the design of the chromatography-chip by

incorporation of elastomeric valves that relieve the pressure accumulated during packing, and are then

closed to restore the column geometry. Adding to these columns an efficient injector geometry which

will also require valves may hamper the fabrication process quality and had complexity to the process.

Other example, also using a PDMS microfluidic structure is this simple microfluidic chip design

for fundamental bioseparation11. The liquid chromatographic column was packed with mesoporous silica

beads. Separation of a dye mixture (fluorescein and rhodamine B) and a biopolymer mixture (dextran

and BSA) was accomplished with good column performance and the chip design was tested. However,

the did not included an on-chip injection system, being handled by off-chip valves and pumping

apparatus.

Finally, the most successful example is the HPLC-chip developed by Agilent Laboratories which

is already available commercially12. This chip includes an enrichment and separation column and a

nanoelectrospray tip all incorporated on-chip for peptide analysis applications. Enrichment and

separation columns were packed with conventional reversed-phase chromatography particles.

19

However, the fabrication process of this chip is impressively more complex and expensive than any

other solution presented so far. Also, the injection system of this device is still considered quite bulky

since a valves system is not fully incorporated.

1.4. Food safety applications

Within the range of relevant analytes for this device, mycotoxins are a particularly important

application since they contaminate the diet of a large proportion of the world’s population, especially in

low-income countries where the contamination levels are the highest13. They represent a serious global

health issue as per most known mycotoxins, like OTA and AFB1, are potential carcinogenic (Group 2B)

and confirmed carcinogenic (Group 1), respectively, for humans14. The fact that such a device can have

a broad range of applications, including critical areas such as food safety, is of upmost importance for

the motivation of this work.

A mycotoxin is a toxic secondary metabolite produced by filamentous fungi, like Aspergillus,

Fusarium and Penicillium. Mycotoxins can appear in the food chain because of fungal infection of crops,

either by being eaten directly by humans or by being used as livestock feed and they can invade crops

directly in the field but also during storage, especially in high humidity and warmer environments.

Aflatoxins, ochratoxins, trichothecenes, zearalenone, fumonisins, tremorgenic toxins, and ergot

alkaloids are the mycotoxins of greatest agroeconomic importance and often more than one mycotoxin

is found in the same contamination site since they are nowadays considered the most dangerous food

contaminants54.

OTA (Figure 10) is produced by fungi of the genera Aspergillus and Penicillium. OTA is a

frequent natural contaminant of many foodstuffs such as cocoa beans, coffee beans, cassava flour,

cereals, fish, peanuts, dried fruits, wine, poultry eggs and milk55.

OTA has a molecular weight of 404 Da and is a weak organic acid (pKa around 4.3 and the

phenol group has a pKa of 7.1) It is soluble in polar organic solvents like alcohols, ketones and

chloroform, but also slightly soluble in water. OTA is a relatively stable molecule under high

Figure 10. Chemical structure of ochratoxin A.

20

temperatures and survives most cooking processes to some extent. Also survives brewing and

winemaking and therefore can be found in a variety of processed consumer food products55.

OTA is considered a nephrotoxic molecule which causes both acute and chronic effects on

kidneys from mammalian species and is also considered genotoxic and teratogenic. Finally, OTA is also

a possible carcinogen, causing renal carcinoma and other cancers in a number of animal species,

although the mechanism for this is uncertain55.

Aflatoxins are poisonous carcinogens that are produced by Aspergillus which grow in soil,

and grains and are regularly found in storage consumables like corn, cotton seed, peanuts, rice, wheat,

and some spices. Animals fed with contaminated food can transmit aflatoxin into products like eggs,

milk derivates, and meat13.

AFB1 has a molecular weight of 312 Da and it is soluble in water and in polar organic

solvents, such as methanol, chloroform, acetone and acetonitrile. AFB1, like OTA, is also a relatively

stable molecule under high temperatures and survives most cooking processes to some extent. AFB1

is considered the most toxic toxin among the other existing variants and it is a known potent genotoxic

and carcinogenic molecule through both damage to DNA and increased oxidative damage13.

A miniaturized chromatographic analytical device where the separation channel constitutes

a packed bed with commercially available beads has a wide range of possible applications, similarly to

conventional HPLC. One of them can be the detection and separation of these toxins for food safety

purposes where fast and cheap analysis could be performed, even with small volume of samples

available. Therefore, this work focus not only on the development but also on the application of the

designed microfluidic for food safety purposes.

Figure 11. Chemical structure of aflatoxin B1.

21

2. Materials and methods

2.1. Microfluidic device fabrication

The fabrication of the microfluidic devices comprised several steps, namely hard mask

fabrication, mold fabrication and replication of PDMS structures. The reagents and materials used for

all these steps and the required equipment are listed in detail in Table 2.

Table 2. Materials and equipment required for microfluidic device fabrication.

2.1.1. Hard mask fabrication

The different design iterations of the microfluidic structure required several hard masks to be

fabricated following the experimental procedure described subsequently in this section and illustrated in

Figure 12. First, a computer assisted design (CAD) of the microfluidic structure was designed using

AutoCAD software for fabrication of the aluminium masks. The substrate used for mask fabrication was

5x5cm glass previously cleaned by washing with acetone, then DI water and lastly immersed in an

Alconox solution for 15 minutes at 65 °C. Completed the washing step, the glass substrate was rinsed

with DI water again (to remove the Alconox solution and remaining dirt) followed by drying with

compressed air. The, a 200 nm aluminium layer was deposited on the clean glass by magnetron

sputtering using a Nordiko 7000 equipment. Afterwards, a positive photoresist (PFR 7790G) layer of 1.5

Step of the process

Reagents/Materials Equipment/Facilities

Hard Mask Fabrication

- Alconox solution, Alconox Inc. (White Plains, NY/USA) - Acetone (99,6%), LabChem Inc. (Zelienople, PA/USA) - isopropanol (IPA), (99,9%), LabChem Inc. (Zelienople, PA/USA) - deionized (DI) water - Glass substrate, Corning Inc, (Corning, NY/USA) - Photoresist PFR 7790G, JSR (Sunnyvale, CA/USA) - Silicon wafer (150 mm diameter), University Wafer (South Boston, MA/USA) - TechniEtch A180 Aluminium etchant, Microchemicals (Ulm, DE)

- AutoCAD software (Autodesk Inc., Mill Valley, CA/USA) - Kerry Ultrasonic Cleaning Bath, Guyson (Skipton, North Yorkshire, UK) - Automatic Dicing SAW DAD-321, Disco Corporation, (Tokyo, JP) - Nordiko 7000 magnetron sputtering system, Nordiko Technical Services Ltd (Havant, Hampshire, UK) - SVG Resist coater and developer track, Silicon Valley Group Inc. (San Jose, CA/USA) - DWL lithograph, Heidelberg Instruments (Heidelberg, DE)

Mold Fabrication

- Silicon substrate, University Wafer (South Boston, MA/USA) - Alconox solution, Alconox Inc. (White Plains, NY/USA) - Acetone (99,6%), LabChem Inc. (Zelienople, PA/USA) - IPA, (99,9%), LabChem Inc. (Zelienople, PA/USA - SU-8 50 photoresist, Microchem Corp. (Newton, MA/USA) - SU8 2015 photoresist, Microchem Corp. (Newton, MA/USA) - AZ 40 XT photoresist, MicroChemicals Corp. (Ulm, Germany) - propylene glycol ether acetate (PGMEA) (99,5%), Sigma-Aldrich (St. Louis, MO/USA) - AZ 400 K developer, MicroChemicals Corp. (Ulm, German) - DI water

- Vertical laminar airflow cabinet, FASTER-BSC-EN (Cornaredo, IT) - Spinner, Laurel Corp. - UVO cleaner 1444AX-220, Jelight Company, Inc. (Irvine, CA/USA) - Hotplate, Stuart (Stafforshine, UK) - UV light (254 nm, 400 W), UV Light Technology Limited (Birmingham, UK) - Stereo microscope, AmScope (Irvine, CA/USA) - Alpha-step 200 profilometer, Tencor Instruments

Fabrication of PDMS

channels

- Sylgard 184 PDMS and curing agent, Dow Corning (Midland, MI/USA) - Micro slides 0.8 mm, Assistent (Sondheim, Germany) - Rounded syringes tips (20 and 18 Gauge), Instech Laboratories, Inc. (Plymouth Meeting, PA/USA)

- Analytical scale d=0.0001g, Scientech (Bradford, MA/USA) - Vacuum desiccator, Bel-Art Products (South Wayre, NJ/USA) - Spin coater, Laurel Corp. - Oven loading model 100-800, Memmert (Schwabach, DE) - Expanded oxygen plasma cleaner PDC-002-CE, Harrick Plasma (Ithaca, NY/USA)

22

μm thick was spin-coated onto the recently deposited aluminium layer. The designs in the AutoCAD file

were then converted and transferred to a DWL laser direct write lithography system (diode laser: 405

nm, blue) that performed a photolithographically transfer of the pattern in the CAD design to the

photoresist. A development step of the photoresist took then place, leaving exposed portions of the

aluminium layer, subsequently removed by a wet chemical etching process with aluminium etchant. To

finish, the remaining photoresist was removed creating a patterned aluminium mask. All the

microfabrication steps previously described were completed under class 100 (100 000 particles over 1

μm, per m3) cleanroom conditions with exception of the photolithography step, which was performed in

class 10 (10 000 particles over 1 μm, per m3) conditions.

Figure 12. Sequence of steps involved in the fabrication of the aluminium hard masks.

2.1.2. Master mold fabrication

The master molds fabrication consisted on several stages of deposition and development of

layers of both different photoresists and heights which were based on previous work56,57. For a clearer

understanding of the experimental procedure, Table 3 shows the runsheet for each master mold and

Figure 13 shows the schematics of different steps involved in the fabrication.

Table 3. Runsheet comprising the sequential steps involved in each mold fabrication process for the different design

iterations, including iterations where two different master molds are necessary (3 and 4).

Fluidic layer mold

Fabrication Steps Iteration 1 Iteration 2 Iteration 3 Iteration 4

Step 1 Silica substrate

cleaning Silica substrate

cleaning Silica substrate

cleaning Silica substrate cleaning

Step 2 20 µm layer 20 µm layer 100 µm layer 20 µm layer

Step 3 100 µm layer 100 µm layer 35 µm layer 100 µm layer

Step 4 Hard bake at 150 ºC

for 15 minutes Hard bake at 150 ºC

for 15 minutes

Final bake by ramping up from 100 to 125 ºC, followed by 5 minutes

baking

35 µm layer

Step 5 - - - Final bake by ramping up

from 100 to 125 ºC, followed by 5 minutes baking

Valves layer mold

Step 1 - - Silica substrate

cleaning Silica substrate cleaning

Step 2 - - 50 µm layer 50 µm layer

- - Hard bake at 150 ºC

for 15 minutes Hard bake at 150 ºC for 15

minutes

23

2.1.2.1. Silicon substrate cleaning

To start the process, a silicon 5x5 cm substrate was cleaned by successive washing steps with

acetone, IPA and DI water to eliminate any residues of photoresist (during the dicing process from the

silicon wafer, photoresist was used to protect the substrate) or other impurities on the surface.

Afterwards, the substrate was immersed in an Alconox solution for 15 minutes at 65 °C, followed by

cleaning with DI water and drying with compressed air. To finish, the substrate was put in a UVO cleaner

for 20 minutes in order to eliminate any lasting organic material on the surface.

2.1.2.2. 20 μm layer

SU-8 2015 (negative photoresist) was spin-coated on top of the substrate for 10 seconds at 500

rpm with an acceleration of 100 rpm/s, followed by a step of 34 seconds at 1700 rpm with an acceleration

of 300 rpm/s, that originated a 20 μm thick layer as measured in the profilometer. Then, a pre-exposure

bake (95 ºC) for 4 minutes was performed in a hot plate with a later step for cooling down of 1 minute.

The hard mask correspondent to the 20 μm design was positioned over the recently deposited resist

layer, carefully placing the aluminium surface facing down so that loss in resolution due to scattering

effects would not happen. The stack was then exposed to UV light during approximately 30 seconds.

One more baking step took place for 5 minutes at 95 °C followed by a 2 minutes’ step for cooling down

to room temperature. Immersion of the substrate in a PGMEA solution was then performed for

development of the pattern exposed on the photoresist with a duration of 2 minutes under manual

agitation. After the development, the substrate was rinsed with IPA and dried with compressed air.

2.1.2.3. 35 μm layer

AZ 40XT (positive photoresist) was spin-coated on top of the substrate for 10 seconds at 500

rpm with an acceleration of 100 rpm/s, followed by a step of 21 seconds, at 2000 rpm with an

acceleration of 1000 rpm/s, that originated a 35 μm thick layer, checked in the profilometer. The spin-

coated photoresist was allowed to settle for 30 minutes to prevent bubbles from being entrapped in the

bulk of the photoresist during the subsequent baking steps. Then, the spin-coated substrate was placed

on a hot plate at 100 ºC, slowly ramping up to 125 ºC, where it continued baking for 5 minutes. The hard

mask correspondent to the 35 μm features was placed on top of the substrate, facing down. The

exposition step took then place during approximately 3 minutes and 30 seconds. Then, a post-exposure

bake was performed at 105 °C for 2 minutes. The mold was then developed by immersion for 10 min in

AZ 400 K developer previously diluted in DI water to a ratio of 1:3, and then washed with DI water.

Finally, a reflow step was performed by ramping up the temperature from 100 °C to 125 °C and then

baking for 5 minutes.

24

Figure 13. Sequence of steps involved in the fabrication of the SU-8 negative photoresist mold (iteration 1, 2 and

valves layer molds) and of SU-8 negative and AZ 40 XT positive photoresist molds (iteration 3 and 4 fluidic layer

molds).

2.1.2.4. 50 μm layer

A SU-8 50 photoresist (negative photoresist) was spin-coated on top of the substrate during 10

seconds at 500 rpm with an acceleration of 100 rpm/s, then followed by 37 seconds at 2300 rpm with

an acceleration of 300 rpm/s. Profilometer measurements were done to confirm the desired height. The

pre-exposure bake step in this stage included sequential baking by heating up to 65 ºC during 3 minutes,

then further heating at 95 ºC for 8 minutes, and finally to cooling down for 1 minute. The hard mask with

the 50 μm features was then manually aligned with the aluminium surface facing down. Exposition to

the UV light was performed during about 25 seconds. Afterwards, a post-exposure bake was done at

65 °C for 1 minute, followed by 7 minutes of heating at 95 °C and, lastly, a 2 minutes’ step for cooling

down. Again, the photoresist layer was developed in PGMEA for about 6 minutes with manual agitation,

washed with IPA, and air dried.

2.1.2.5. 100 μm layer

SU-8 50 photoresist was spin-coated on top of the substrate during 10 seconds at 500 rpm with

an acceleration of 100 rpm/s, followed by 30 seconds at 1000 rpm with an acceleration of 300 rpm/s.

25

The pre-exposure bake step in this stage included sequential baking by heating up to 65 ºC during 10

minutes, then further heating at 95 ºC for 30 minutes, and finally cooling down for 1 minute. The hard

mask with the 100 μm features was then manually aligned with the aluminium surface facing down.

Exposition to the UV light was performed during about 70 seconds. Afterwards, a post-exposure bake

was done at 65 °C for 1 minute, followed by 10 minutes of heating at 95 °C and, lastly, a 2 minutes’ step

for cooling down. Again, the photoresist layer was developed in PGMEA for about 10 minutes with

manual agitation, washed with IPA, and dried.

2.1.3. Fabrication of PDMS channels

The finished master molds were then used for the fabrication of the PDMS channels. For the

microfluidic structure iterations that included valves, besides different master molds they also included

different fabrication processes for the PDMS channels which are described in this section along with the

standard fabrication process and were based on recent published work from the group57. A scheme

highlighting the major differences in the process is shown in Figure 14.

Figure 14. Sequence of steps involved in the fabrication of the PDMS channels for the different iterations, using

distinct fabrication methods: soft lithography with irreversible sealing done by plasma treatment (iteration 1 and 2);

multilayer soft lithography and sealing using an adhesive PDMS layer (iteration 3 and 4A) or plasma treatment

(iteration 4B).

26

2.1.3.1. Process for iteration 1, 2 and microcolumns

The first step was to fix the mold in a Petri dish with the patterns facing up with little pieces of

tape. PDMS elastomer preparation was done by the following experimental procedure: a curing agent

and PDMS mixture was prepared in a plastic cup in a 10:1 ratio, followed by a degasification step for 30

minutes in a desicator. After this, the mixture was poured on top of the mold (inside the Petri dish) until

it fills about half of the height of the latter. The PDMS was cured for 90 minutes at 70 ºC for curing. The

cured PDMS was afterwards cut by means of a scalpel and peeled off from the mold with appropriate

tweezers. Holes were punched on the structures with rounded 20 and 18 Gauge needles for the outlets

and inlets, respectively. A 500 μm thick PDMS membrane was prepared with the purpose of sealing the

PDMS structures, by spin-coating PDMS from the mixture previously prepared on top of a silicon wafer

at 250 rpm for 25 seconds with an acceleration of 100 rpm/s. A baking step was performed as described

above and cut in parts with an appropriate size. Both the structures and the membranes were taken into

an oxygen plasma cleaner in order to seal the two against each other, immediately after plasma

treatment.

2.1.3.2. Process for iteration 3 and 4A

The procedure for PDMS preparation was the same as described in the previous section, in

addition that PDMS on a ratio of 20:1 was also prepared. The PDMS preparation of 10:1 ratio was

poured on top of the fluidic layer mold and baked at 70°C for 50 min while the PDMS preparation of 20:1

ratio was spin-coated on top of the valves layer mold for 25 seconds at 300 (400 μm thickness), 400 or

500 rpm with an acceleration of 100 rpm/s and baked at 70 ºC for 40 minutes. After the half-curing

process, the PDMS was taken from the mold and inlets/outlets were punched using 20 Gauge blunt

syringe tips. The fluidic layer was then aligned on top of the valves layer and the stack was baked

together at 70 °C for 90 minutes. Completed the curing process, the structures were cut and

subsequently taken from the mold with both layers irreversibly sealed against each other. The inlets

corresponding to the valves channels were punched using again 20 Gauge blunt syringe tips. Instead

of an oxidation treatment, sealing was achieved by gently wet the PDMS stack on a PDMS adhesive

layer obtained by spin-coating PDMS 10:1 on top of a coverglass (1 mm thick) for 15 seconds at 500

rpm, followed by 30 seconds at 3000 rpm and, finally, 4 minutes at 6000 rpm. Then, the stack was

sealed against a 100 μm thick glass slide in which PDMS was previously spin-coated for 5 seconds at

500 rpm, followed by 20 seconds at 4200 rpm, resulting in a PDMS-PDMS sealing after a final bake at

70 °C for 90 minutes.

2.1.3.3. Process for iteration 4B

Both the PDMS preparation and casting was done as described in the previous section only

varying the PDMS curing time from 50 (fluidic layer) and 40 (valves layer) minutes to 90 minutes for

27

each layer. The two different PDMS layers were taken into an oxygen plasma cleaner in order to seal

the two against each other, immediately after plasma treatment, creating a PDMS stack that was further

sealed against a 1 mm-thick glass slide in which PDMS was previously spin-coated for 5 seconds at

500 rpm, followed by 20 seconds at 4200 rpm, resulting in a PDMS-PDMS sealing using again the

oxygen plasma cleaner. Between each plasma treatment, the structures were allowed to stabilize a

minimum of two hours before being used in order to increase the robustness of the bonding and avoid

eventual sealing of the valves channels to the surrounding PDMS, upon actuation with the compressed

air.

2.2. Microfluidic device manipulation

The fabrication of the microfluidic devices comprised several steps, namely separation channel

packing and separation and elution studies. The reagents and materials used for all these steps and the

required equipment are listed in detail in Table 4.

Table 4. Materials and equipment required for microfluidic device manipulation.

2.2.1. Separation channel packing

Different beads were used for each experiment were used throughout the experimental work as

well different microfluidic structure configurations which demanded both tailored bead preparation

protocols and packing methodologies. Both subjects are further described in this section.

Step of the process

Reagents/Materials Equipment/Facilities

Separation channel packing

- Insulin syringe 1 mL U-100 Luer-Lock, Codan (Lensahn, DE) - Tubing couplers SC20/15, Instech Laboratories, Inc. (Plymouth Meeting, PA/USA) - Polyethylene tubing (BTPE-90), Instech Laboratories, Inc. (Plymouth Meeting, PA/USA) - Milli-Q water 18 MΩ CM, Millipore - Polyethylene glycol (PEG) 8000 MW, Sigma-Aldrich (USA) - Ethanol (96%), Sigma-Aldrich (USA) - CM-Sepharose Fast Flow Beads 90 μm, GE Healthcare Life Sciences 17-0719-05, (USA) - Spherical C18 bonded flash silica beads 45-75 μm, 97727-U Supelco Analytical (Bellefonte, PA) - Spherical flash silica beads 45-75 μm, 97728-U Supelco Analytical (Bellefonte, PA) - Triton X-100, Sigma-Aldrich (USA)

- Syringe pump NE-1002X, New Era Pump Systems, Inc. (Farmingdale, NY/USA) - 170 mesh cell strainer Alfa Aesar, ThermoFisher (Karlsruhe, GE) - Leica DMLM Microscope, Leica Microsystems (Wetzlar, DE)

Separation and elution

studies

- Insulin syringe 1 mL U-100 Luer-Lock, Codan (Lensahn, DE) - Tubing couplers SC20/15, Instech Laboratories, Inc. (Plymouth Meeting, PA/USA) - Polyethylene tubing (BTPE-90), Instech Laboratories, Inc. (Plymouth Meeting, PA/USA) - Milli-Q water 18 MΩ CM, Millipore - Methylene blue stain, Sigma-Aldrich (USA) - Yellow food colouring E104, E122, E260, 5160414 Globo, (UK) - Phosphate pH 7.5 1M NaCl at 25 ºC solution, Sigma-Aldrich (USA) - Phosphate pH 2.5 2M NaCl at 25 ºC solution, Sigma-Aldrich (USA) - Milli-Q water (18 MΩ CM), Millipore - HCL 1M solution, Sigma-Aldrich (USA) - FITC F427 (98%), Sigma Aldrich (Switzerland) - Aflatoxin B1 from Aspergillus flavus, Sigma-Aldrich (USA) - OTA from Aspergillus ochraceus, Sigma-Aldrich (USA) - Acetonitrile solution (99,8%), Sigma-Aldrich (USA) - Methanol solution (99.9%), Sigma-Aldrich (USA) - white polystyrene 96 well plates, Corning (NY/USA)

- Syringe pump NE-1002X, New Era Pump Systems, Inc. (Farmingdale, NY/USA) - Fluorescence spectrometer Varian, Cary Eclipse - Leica DMLM Microscope, Leica Microsystems (Wetzlar, DE) - ImageJ software (USA)

28

2.2.1.1. Beads preparation protocols

Commercially available beads were supplied in an ethanol solution (20% v/v). For CM-

Sepharose Fast Flow Beads, the first step was sieving the beads, due to their large particle size

dispersion (45 μm-165 μm), using a 170 mesh. The next step was to homogenize the bead suspension

and transfer a certain volume of the bead stock solution into a PEG 8000 20% (w/w) solution in a

proportion of 1:4. PEG 20% was used to ensure beads were homogeneously dispersed and to prevent

settling, which would cause clogging of the microchannel with clumps of beads before an adequate

packing being achieved. For the C18 beads a different protocol was followed to prevent beads

aggregation in solution due to their high hydrophobicity and assure an easy packing inside the

separation channel. From a commercial dry stock of C18 beads, approximately 10 mg were weighed in

an analytical balance and further mixed in 600 μL of previously prepared PEG 20% and 2 μL of Triton

X-100. The prepared solution was mixed continuously for about 30 seconds in the vortex, repeating the

step 2-3 times until complete homogenization was achieved. For silica beads and different beads used

throughout the experimental work, when not specified otherwise, the following preparation protocol was

used: beads were always prepared in stock solution of 20% ethanol and further homogenised in vortex

to then collect a certain volume of stock solution that was added to a PEG 8000 20% (w/w) solution in

a proportion of 1:4 and finally used in the packing and separation experiments.

2.2.1.2. Packing method

In order to pack the beads rapidly and efficiently, three different packing methodologies were

used throughout the work: either a syringe pump on a pushing or pulling mode or a compressed air

packing. To pack the separation channel using the pushing mode (positive pressure applied) the flow

applied to fill the separation channel varied in a range of 10-20 μL/min by action of a syringe pump.

Beads suspension entered the column in a 20 Gauge inlet via a capillary tubing pre-filled with water and

beads solution and connected to the structure using a metal coupler. Caution measures were taken in

order to the capillary tube to be free of air bubbles. To avoid channel clumping during filling, continuous

gentle peristaltic movements were performed on the channels structure with a tweezers, again to avoid

clumping on the channels. Alternatively, using the syringe pump in pulling mode the flow was driven by

applying a negative pressure at the outlet, via capillary tubing pre-filled with water and connected to the

microfluidic structure with a metal coupler inserted into the outlet. A pipette tip containing the bead

suspension was inserted into the inlet and the packing was performed at a flow rate between 10 to 15

µL/min. Finally, the compressed air packing consisted in filling the separation channel with beads

solution using capillary tubing connected to compressed air lines and subjected to a pressure of 0.15

MPa. The compressed air lines were switched on or off via solenoid valves connected to a compressed

air supply that were controlled through a printed circuit board (PCB) supplied by a 24 V power source

(Figure 15).

29

Completed the filling of the channels, inlets and outlets were sealed using closed metal plugs

to avoid solution loss and the structure was stored on the fridge immersed in DI water, approximately at

4 ºC for further uses.

2.2.2. Separation and elution studies

Experimental work in order to validate the functionality of the microfluidic structures and assess

the potential application on food safety analysis was done and the methodologies are described below

in this section.

2.2.2.1. Food colouring separation (proof-of-concept)

Filtered CM-Sepharose Fast Flow bead suspension was prepared and packed inside the

separation channel as described in section 2.2.1.1 and 2.2.1.2. Then, Milli-Q water was flowed during

about 10 minutes at a flow rate of 15 µL/min in order to wash the separation channel from the

components of the beads packing solution. A green solution resulting from the mixture of methylene

blue and yellow food colouring in a proportion of 1:1 was prepared. A volume of about 10 μL of green

solution was used to insert in a capillary tubing pre-filled with water and connected to the structure using

a metal coupler inserted on the cross-shaped injector inlets. To insert a small plug in the channels, using

a flow rate of 3 μL/min, until near 1/3 of the total volume of the green solution entered the channel.

Afterwards, Milli-Q water was used as mobile phase, with a constant flow rate of 8 μL/min in pushing

mode (positive pressure at the inlet) by action of a syringe pump. Elution of the methylene blue retained

on the beads was tested with 3 different buffers/solutions: (i) phosphate buffer at pH 7.5 containing 1 M

NaCl, (ii) phosphate buffer at pH 7.5 containing 2 M of NaCl, (iii) 1 M HCl solution. The flow rate applied

Figure 15. Experimental setup for compressed air packing and valves actuation. (A) Switch buttons for on/off

actuation on the valves. (B) and (C) PCB for controlling of valves actuation and respective power source. (D)

Syringe pump. (E) Microfluidic device connected to the system by capillary tubing.

30

for elution steps was always a constant value of 13 μL/min during at least 20 minutes, in order to

completely remove the methylene blue retained on the beads. After each elution assay, the column was

washed with Milli-Q, during about 10 minutes at 15 µL/min, in order to remove any residual

buffer/solution flowed previously.

2.2.2.2. OTA and AFB1 microchannel adsorption and elution

OTA and AFB1 stock solutions at 100 µg/mL were used to prepare diluted solutions in Milli-Q

water of 10, 6, 3 and 1 µg/mL each. Adsorption was performed in already packed microchannels

designed in previous work56 packed with C18 beads. Milli-Q water was flowed during about 5 minutes

at a flow rate of 15 µL/min in order to wash out the components of the bead packing solution. A pipette

tip containing 10 µL of 6 µg/mL of each toxin solution was inserted in the inlet and the solution was

flowed at a flow rate of 15 µL/min, by applying a negative pressure at the outlet. Acetonitrile and

methanol solutions of 100, 75, 50, and 25% (v/v) in Milli-Q water were prepared and tested with respect

to the ability of eluting the adsorbed toxins elution. Acetonitrile and methanol solutions were flowed

inside the microchannel by applying a positive pressure (pushing) at the inlet at a flow rate of 5 µL/min

for 10 minutes. For image acquisition during elution, auto fluorescence of the toxins was monitored

every 30 seconds using a DFC300FX digital camera coupled to a Leica DMLM microscope with UV light

excitation filter. The UV light excitation filter has a band pass illumination path at wavelengths between

355 and 425nm and a long pass observation path above 470nm. Images were taken with an exposure

time of 1 second and 1x gain and the average fluorescence intensity was measured using ImageJ

software.

To assess the solvent effect in the fluorescence of the toxins measured with the miscroscope,

solutions of 6 µg/mL of both OTA and AFB1 in 100, 75, 50, 25%, 0% (v/v) of either acetonitrile or

methanol in Milli-Q water were prepared and placed on 96 well plates. Measurements were performed

in a fluorescence spectrometer with an excitation and emission wavelengths of 333nm and 460 nm,

respectively, and excitation/emission slits of 5 nm.

2.2.2.3. OTA and AFB1 cross-channel injection and elution

In order to study the feasibility of AFB1 elution within a packed column with a cross-channel

injector (iteration 2) and also the separation of AFB1 and OTA within the same column, the fluorescence

of the toxins was followed in the packed channel. C18 bead suspension was prepared and packed inside

the separation channel as described in sections 2.2.1.1 and 2.2.1.2. Milli-Q water was flowed during

about 10 minutes at a flow rate of 15 µL/min in order to wash out the components of the bead packing

solution. A pipette tip containing 60 µL of 6 µg/mL of AFB1 solution (for the elution) or 60 µL of 6 µg/mL

of AFB1 + OTA solution (for the separation) was inserted in the inlet and the solution was flowed at a

flow rate of 15 µL/min, by applying a negative pressure at the outlet of the cross-channel injector.

31

Acetonitrile 100% v/v (for the elution) and 25% v/v (for the separation) in Milli-Q water solutions were

flowed inside the microchannel by applying a positive pressure (pushing) at the inlet at a flow rate of 2

µL/min for approximately 10 minutes. For image acquisition during elution, auto fluorescence of the

toxins was monitored using a DFC300FX digital camera coupled to a Leica DMLM microscope with UV

light excitation filter. The UV light excitation filter has a band pass illumination path at wavelengths

between 355 and 425nm and a long pass observation path above 470nm. Images were taken with an

exposure time of 1 second and 1x gain.

2.2.2.4. Plug optimization

In order to study plug formation and progression inside a microfluidic structure with a cross-

shaped injector actuated by pneumatic valves (iteration 4B), plug optimization assays were performed.

An ethanol 96% (v/v) solution was flowed by applying a positive pressure (pushing) using a syringe

pump at a flow rate of 15 µL/min during 20 minutes. All inlets should be closed using a pipette tip filled

with liquid to avoid bubble trapping inside the injector and separation channel. Microfluidic structures

with and without packed silica beads were used for assessment of the effect of beads in plug

progression. FITC at 1 mg/mL solution in ethanol 96% was prepared and a volume of 5 µL was loaded

onto the cross-shaped injector by pulling at 15 µL/min and subsequent actuation of the valves. The

ethanol 96% solution was flowed inside the channel at a flow rate of 0.5 µL/min by pushing with one or

more syringe pumps. For image acquisition, during operation, the fluorescence of the FITC solution was

monitored using a DFC300FX digital camera coupled to a Leica DMLM microscope with a blue light

excitation filter. The blue light excitation filter has a band pass illumination path at wavelengths between

450 and 490 nm and a long pass observation path above 515nm. Images were acquired with an

exposure time of 800 milliseconds and 3x gain.

32

3. Results and Discussion

3.1. Microfluidic device design considerations

The first step into the development of the chromatographic microfluidic device was the

conception of the design of both the separation channel and later, the injector geometry. Many factors

must be taken into consideration, from the aspect ratio height/width to particle size to the channel length

and beads trapping within the column46. Conventional columns typically comprise a glass, plastic or

metal cylinder with a packed stationary phase, and various plumbing arrangements to allow introduction

of the sample and collection of fractions with minimal mixing, peak distortion, dilution and sample loss.

In a HPLC, the most common columns have a diameter around 4.6 mm and a length from 10 to 15 cm,

packed with 5 µm beads, although many other sizes can be found16. In microfluidics, microchannels

(analogous to the column) are typically around 100-150 μm in height and can have varied widths and

lengths, only restrained by the scale of wafers and lithography processes.

In Figure 16, the comparison between the two initial CAD iterations for the design of the

microfluidic device are shown. Both iterations have the same separation channel height and width, 100

and 400 µm, respectively. The ratio of channel width (or height, depending on the characteristic length)

to particle size must be considered to avoid the known “wall effect” where quality/density of packing or

flow behaviour will be severely different from the bulk channel, which is very common in microfluidic

devices where aspect ratios of width to particle size are usually around 1046,47. In the case of our

separation channel dimensions, the aspect ratio will be of 4.4 and 6.9 for the beads with mean diameters

of 90 and 58 µm, respectively, that were used throughout the experimental work. For values much lower

than 10 the wall and the bulk region become very similar, therefore such effect loses its relevance.

However, the aspect ratios values are still very close to ignore the effect on quality/density of packing

and flow behaviour which will diminish the column efficiency. To overcome this limitation, several

strategies could be employed on the design: (i) decrease the width of the channel, (ii) decrease the

particle size or (iii) increase the length of the channel. Decreasing the width of the channel would lead

to values much lower than 10 and therefore the effect would not be significant, as previously mentioned.

However, this would also reduce the surface area available for interaction with the analytes and mobile

phase since the number of beads packed would severely decrease. On the other hand, decreasing the

particle size would increase the value of the aspect ratio to 80 if, for example, 5 µm beads were used

and therefore the wall region would only constitute a small part of the channel, reducing the impact of

this effect. This has been a strategy used both in conventional and on-chip HPLC44, however with the

consequence of increased pressure demand on the devices. Also, due to the bead trapping feature

being based on the different heights of the channel and outlet, such a small outlet cannot be fabricated

with the current instrumentation and other bead trapping strategies would have to be employed. Finally,

there is also the opportunity of increasing the length of the channel. Based on theoretical knowledge on

chromatography, it is well known that an increase in length of the column will provide a higher column

33

efficiency value (N) as shown previously in Equation 2 (𝑁 = 𝐿 𝐻⁄ ), where N is the number of theoretical

plates, L the length of the column and H the plate height. This would compensate that loss in separation

efficiency due to the wall effect.

Therefore, an initial length of 30 cm was chosen for the separation channel, being possible by

the serpentine design geometry chosen (Figure 16, 1-B) that maximizes the area available within the

5x5 cm mold. However, due to restrictions in the applied pressures (see section 3.3) during beads

packing, in iteration 2 the separation channel length had to be reduced on more than 50% (12 cm) in

order to be possible to pack the whole channel. The impact of this reduction will be further addressed in

the next sections.

Figure 16. Comparison between the first two CAD iterations created using AutoCAD. (1-A) corresponds to a close-

up representation of the bead trapping feature near the outlet where the channel has two different heights: 100 and

20 µm. (1-B) corresponds to a microscopic image of the master mold fabricated with the serpentine separation

channel with a total length of 30 cm. (1-C) corresponds to the microscopic image of the master mold where the

bead trapping can be seen. (2-A, B) highlight the simple cross-channel injector representation introduced into the

design near the inlet and a microscopic image of the injector on the master mold, with the two channels of different

heights (100 and 20 µm). (2-C) Microscopic image of the bead trapping feature and serpentine separation channel

with a total length of 12 cm.

34

Another important aspect common to both designs is the bead trapping feature which plays with

the difference in the heights of the channels to entrap the beads (Figure 16, 1-A, C). The main purpose

was to retain the bead packed bed by creating a outlet channel which has a height much smaller than

the average bead size, working as a barrier for beads. Since the beads to be used in further experimental

work had average particle size of no more than 100 μm, channel heights were defined as 100 and 20

μm for the larger and smaller channel, respectively. The trapping feature proved to be efficient as it can

be seen further ahead in section 3.3, also probably allowing better column performance in terms of band

broadening since it does not require so much dead volume and disturbance in flow as other solutions51.

The spacing between channels was always above 400 µm to prevent problems in the microfabrication

process (for example, detaching of the photoresist layers from the substrate due to extreme small

features) which proven to be enough for robust fabrication as seen in the microscopic images of Figure

16.

One difference to note between the two iterations are the inlets and outlets, at two levels: first,

both inlet and outlets were reduced in diameter (Figure 16, iteration 2) due to collapsing of the inlet on

the PDMS membrane that was used to seal the structures; secondly, the formation of clogs at the inlet

when inserting the bead suspension led to alterations in the inlet shape so that beads flow would be

facilitated at the beginning of the separation channel.

Finally, the single inlet geometry was replaced by a cross-channel injector in iteration 2 (Figure

16, 2-A, B) to allow smaller and well-defined sample plugs. The importance of the injector as a

component in a microfluidic separation system arises from the fact that it defines the shape and quantity

of analyte that will be used for separation and analysis. The cross-channel injector is a simple

intersection of two channels that also have different heights, similarly to the beads trapping feature, to

ensure beads are kept within the separation channel. It is operated by first loading the beads solution

into the central inlet (oval-shaped), followed by insertion of sample solution on the perpendicular channel

inlet (rounded-one). A defined sample plug will be originated and travel through the separation channel

by loading the mobile phase on the central inlet. However, this cross-channel geometry controlled off-

chip with syringe pumps originated still an uncontrolled and irreproducible loading of sample and if not

carefully handled by an experienced user, the plug created could be as large as one defined without a

injector. These problems arouse mainly due to the rapid plug axial diffusion with time and pressures

differences created from insertion and removal of metal plugs to control fluid flow.

Currently, attempts to adapt pressure-driven separations to a microfluidic device requires the

use of conventional, off-chip sample definition or very complex on-chip injectors12. The large volume

and dead time introduced by an off-chip injection hampers the ability of a microfluidic separation to

deliver rapid and sensitive analyses. A new technique developed by Unger et al. designated multilayer

soft lithography39,40 allows the fabrication of integrated pumps and valves using elastomeric material,

such as PDMS. In Figure 17, a comparison between iterations 3 and 4 is shown where the major

35

difference between the two designs is the injector geometry and location and how the pneumatic

actuated valves, based on the work of Unger et al., were integrated within the microfluidic device.

In iteration 3, the cross-channel injector geometry was maintained, still using the height

difference feature to entrap the beads within the separation channel. However, the inlets for beads

insertion and mobile phase flow were separated (Figure 17, 3-C) so that once totally packed, the

separation channel would be blocked to prevent the unwanted movement of the packed bed and sample

plug when pressure differences were created from the insertion/removal of the metal plugs to control

fluid flow. Valves were incorporated underneath the cross-channel injector inlet and outlets and on the

separation channel outlet with an actuation area of 200 x 200 µm. Controlling the fluid loading and flow

using integrated valves provided the generation of well-defined and controlled plugs and injections.

However, elastomers deform in response to pressure, creating fluidic capacitance that allied to the

cross-channel geometry caused the same problems related to pressure differences from the injector of

Figure 17. Comparison between the third and fourth CAD iterations created using AutoCAD. (3-A) corresponds to a

close-up representation of the cross-channel injector with integrated pneumatically actuated valves. (3-B) corresponds

to a microscopic image of the valves mold fabricated showing a single valve channel. (3-C) corresponds to the

microscopic image of the master mold where the separation channel and cross-channel injector can be seen. (4-A,

C) A close-up representation and microscopy image, respectively, highlights the connection channel (height: 20 µm)

designed between the new cross-channel injector (height: 35 µm) and the separation channel (height: 100 µm) which

entraps beads within the separation channel. (4-B) Microscopic image of the new incorporated cross-channel injector

with integrated pneumatically actuated valves.

36

iteration 2 as seen in Figure 18-A where beads are invading the cross-channel injector and leaving the

packed bed.

To overcome the difficulties encountered in the cross-channel geometry, the injector was

detached from the separation channel to not intersect with the packed beads bed and interfere with

beads packing and plug formation. In iteration 4, the injector consists of two perpendicular channels

(Figure 17, 4-B) with the same height (35 µm) that were pneumatically actuated with the valves channels

underneath (actuation area of 400 x 400 µm). From the microscopic image it is possible to note that the

injector channels on the mold have rounded edges, a requirement for every channel that will be actuated

with the pneumatic valves, since they can only fully close if the channel has a round cross-section. A

connection channel (Figure 17, A-C) of 20 µm height was designed to connect the injector and

separation channel and prevent beads invasion to the injector, using the same principle of the beads

trapping feature. In Figure 18-B, it is possible to see that although the beads can invade the connection

channel at packing pressures higher than 0.2 MPa, they do not reach the cross-channel injector. Finally,

the valve actuating on the outlet of the separation channel was removed in iteration 4 since the

resistance to fluid flow created from the packed bed was so high that any pressure difference effect that

could compromise plug injection was not relevant.

Figure 18. Comparison between the two injectors from iteration 3 and iteration 4. (A) Microscopic image of the

cross-channel injector and separation channel from iteration 3, packed with silica beads. (B) Microscopic image of

the cross-channel injector, connection channels and separation channel from iteration 4, packed with silica beads.

37

3.2. Fabrication process considerations

Besides improving the design from iteration to iteration, many improvements were also needed

regarding the fabrication process itself, especially related to the valves optimization and the sealing of

the microfluidic devices comprising iteration 3 and 4.

3.2.1. Valves thickness optimization

The working principle of the pneumatic valves is based on the deflection of a PDMS membrane

(valves layer) beneath the cross-channel injector, caused by the injection of compressed air in the valve

channels filled with water. It is important to highlight again that the actuated channels need to be

fabricated with a round cross-section, to ensure their complete closure. Based on previous work from

the group57, the needed PDMS valves’ layer thickness for complete closure of the channels by the valves

at 0.1 MPa was 400 µm (300 rpm spin-coating speed). Therefore, the valves layer was first fabricated

using this spin-coating speed. However, as seen from Figure 19-E, the actuation of this valve at the

same pressure (0.1 MPa) in the microfluidic device was not enough to fully close the channel. The

different results can be caused by the difference in the actuation area of the valve or by the pressure

that device accumulates, which is much higher than the device from the reported work due to more

complexity in the design and a higher separation channel length. Applying higher pressures to close the

Figure 19. Microscopic images PDMS microfluidic structures (iteration 3) comprising valves of different thicknesses

originated by different spin-coating speeds, and its actuation. (A, B and C) are microscopic images of valve channels

spin-coated at 300, 400 and 500 rpm originating decreasing valves layer thickness. (D and E) are microscopic

images of the actuation of valve channels at P = 0 and 0.1 MPa, respectively, for a 300 rpm spin-coated layer. (F)

is a microscopic image pf the actuation of valve channel at P = 0.1 MPa for a 400 rpm spin-coated layer. Green

food colouring was flowed through the injector channel for better visualization of valves actuation.

38

valves eventually led to rupture of the microfluidic structure, consequently decreasing the valves layer

thickness by increasing the spin-coating speed was the followed strategy to overcome this limitation.

Spin-coatings speeds of 400 and 500 rpm were further tested as seen in Figure 19-B and C, respectively.

At 500 rpm, the valves layer eventually became too thin causing the valve channels distortion due to the

pressure applied when sealing the several layers of PDMS into one microfluidic structure. On the other

hand, the 400 rpm spin-coating speed allowed for a good fabrication quality without channel distortion

(Figure 19-B) and a P = 0.1 MPa was enough to fully close the channel, as seen in Figure 19-F.

Therefore, the 400 rpm spin-coating speed was the chosen one for further fabrications using either

iteration 3 or 4 microfluidic structures.

3.2.2. Microfluidic structure sealing optimization

Sealing of the PDMS microfluidic structures is an important step during fabrication, and many

techniques have been employed in order to achieve really robust bonding onto PDMS membranes58 or

glass59. During the experimental work done, both the conventional plasma cleaning strategy and

bonding with PDMS glue and half-curing strategy were employed on the different iterations (Figure 20).

Plasma cleaner treatment (oxidation) creates silanol groups by converting –O2Si(CH3)2 to –

O2Si(OH)2. The surfaces undergo an irreversible bonding because of the siloxane covalent bonds (Si–

O–Si) that are formed during condensation reactions. The oxidative effect in PDMS is temporary owing

to the diffusion of oligomers from the bulk to the surface. Hence, the bonding step must be performed

Figure 20. Schematics illustrating the two strategies used for sealing of the microfluidics structures comprising

valves (iteration 3 and 4), namely bonding through half-cure and PDMS glue or bonding with conventional plasma

cleaner treatment.

39

within one minute after the oxidation treatment. This fact hinders the large-scale integration and steps

of alignment for complex devices.

Plasma cleaner treatment technique was employed for bonding glass or PDMS membranes to

microfluidic structures that comprised the iteration 2 design or the microcolumns due to its simplicity,

robustness and easy execution. However, for iterations 3 and 4 that included a multilayer microfluidic

structure, bonding between layers promised to be more complex. Based on fabrication troubleshooting

reports from previous work in the group, bonding the valves thin layer to the fluidic layer using plasma

cleaner treatment lead to collapsing of the valves during the sealing process (when both surfaces were

hydrophilic), causing irreversible bonding of the valves to the fluidic layer and consequently, the valves

membrane did not respond to actuation. Alternatively, MLS technique offers a bonding without oxidative

treatment, based on the capability to bond multiple patterned layers through half-curing of each layer

and baking them again stacked together to complete the curing process. The principle is to have two

opposite (the ratio of element A and B in the elastomer mixture is substantially unalike) layers in contact

with each other since reactive molecules remain at the interface between the layers and can be further

cured to produce a sealing. Finally, for iteration 3 and 4, besides the interlayer bonding, the final stack

had to be sealed against a glass surface which conferred more robustness to the final structure. Since

MLS cannot be employed to glass surfaces, a thin glass with a half-cured PDMS layer was used and

bonding was performed using PDMS glue (Figure 20). The PDMS glue method consists in wetting the

surface of the microfluidic stack onto a uncured 10:1 PDMS layer spin-coated on a glass. This adhesive

layer created on the microfluidic stack will then be aligned on top of the thin glass with the half-cured

PDMS layer and baked together to finish the curing process.

However, bonding the several layers and glass using MLS and PDMS glue offers low robustness

of the microfluidic structure, withstanding badly high pressures that lead to rupture and leaks of the

structure during operation. Therefore, the bonding using plasma cleaner treatment for iteration 4 was

also employed and compared with the alternative, as seen in Figure 20. Surprisingly, the bonding with

plasma treatment did not interfere the actuation of the valves, as seen in section 3.6, where the

microfluidic structures sealed through plasma treatment are being used without impediments.

Additionally, the structures fabricated with this process (iteration 4B) were more robust when compared

to iterations 3 and 4A and did not had leaks or ruptures as often as the previous ones, as expected. This

result may be explained by the fact that actuation areas of the valves reported on the work were larger

(700 x 500 µm), and therefore more easily collapsed during the sealing process unlike the ones used

on this microfluidic device (400 x 400 µm.)

3.3. Separation channel packing

One of the most important steps throughout the development of the microfluidic device was the

packing of the separation channel with beads. From the bead packing solution to, most importantly, the

packing methodology employed, many factors had to improved.

40

Table 5. The different packing solutions required for each beads type.

Beads type Packing solution

CM-Sepharose, 90 µm Beads suspension of ethanol 20% (v/v) on a PEG

8000 20% (w/w) solution in a proportion 1:4 Silica beads, 58 µm

C18 bonded silica beads, 58 µm 10 mg of dry C18 beads in a PEG 8000 20%

(w/w) solution + 2 µL of Triton X-100

In Table 5, the different beads types used throughout this work is listed and the corresponding

packing solutions used for each one. PEG 20% was used to ensure beads were homogeneously

dispersed and to prevent settling, which would cause clogging of the separation channel with clumps of

beads before an adequate packing being achieved. Due to its hydrophilic nature, PEG was a good

dispersant for silica and CM-Sepharose beads which are also more hydrophilic. However, a different

packing solution had to be used for C18 bonded silica beads since they were very hydrophobic and

would create aggregates, aggravating severely the packing of the beads into the channel. Additionally

to PEG 20%, Triton X-100 was added to the packing solution for C18 beads. Triton X-100 is a non-ionic

detergent, considered a mild surfactant with a hydrophobic character that helps to homogeneously

disperse beads and avoid aggregation to PDMS channel walls. Besides the packing solution, one factor

to consider during the beads preparation procedures was the particle size distribution of the beads. CM-

Sepharose beads have a particle size distribution of 45-165 µm, although the mean size is of 90 µm

which led to clogging problems in the channels. The maximum height of the channels was 100 μm,

being inevitable that beads got stuck in the channel, not allowing correct column packing. Therefore,

beads sieving was necessary to be included in the beads preparation procedure for CM-Sepharose

where they were sieved on 170 mesh that excluded every bead that was above 90 µm in size. After

sieving, beads were successfully packed within the separation channel as seen in 3.4, where

experiments were done using a microfluidic structure packed with sieved CM-Sepharose beads.

Three main strategies were employed to pack the beads within the separation channel as

illustrated in Figure 21 (centre). The first one was to apply a negative pressure at the outlet of the

separation channel (packing by pulling) at a constant flow rate which would flow the beads from pipette

tip into the channel. The second one was to directly flow the beads solution by applying a positive

pressure (packing by pushing) at the inlet of the separation channel, maintaining the flow rate constant.

Finally, the third strategy was to pack the beads by directly flowing the beads solution into the inlet of

the separation channel at a constant pressure applied by a compressed air line. The pressure applied

to flow the beads could be controlled up to 0.3 MPa. The first two strategies are based on packing by

maintaining the flow rate constant, unlike the latter, which packs the separation channel by controlling

the pressure applied. This difference was very important to be able to pack the very long separation

channels used in this microfluidic device.

41

The “packing by pulling” strategy would be preferential since the use of a negative pressure

provides a gradual increase in liquid flow velocity with minor distortion of the PDMS channel or

deformation of the beads, contrary to what is observed when a positive pressure was used, where there

is also a higher likelihood of trapping more air bubbles. However, due to the extended length of the

separation channel, a high pressure difference is required to enable the flowing of beads solution within

the channel. Therefore, it was not possible to overcome this limitation and packing by this method was

disregarded.

Regarding the “packing by pushing” strategy, although it has some of the previously mentioned

drawbacks, it can easily overcome the limitation imposed by the “packing by pulling” strategy, and

several microfluidic structures were packed within 40 minutes using sieved CM-Sepharose beads further

used for the experiments in section 3.4. Packing separation channels longer than 12 cm with this method

proved to be impossible with any type of beads due to the high pressure requirements that the

microfluidic structure could not withstand, provoking leaking and rupture, mainly at the cross-channel

injector area. This phenomenon is easily deduced from equation 6:

Figure 21. Schematics comprising a timeframe of the beads trapping feature with packed silica beads (left); a

comparison between the different packing methods used throughout the experimental work (centre); and a set of

microscopic images of a fully packed 12 cm length separation channel with C18 bonded silica beads (right, A-D).

42

∆𝑃 ~ 𝐿2𝜇

𝑏2𝑡𝑅

where the increase of the separation channel length is directly correlated to the increase of the

pressure requirements for the structure.

However, this strategy revealed to not be satisfying when using C18 bonded silica beads, which

due to their hydrophobic nature offered a lot more resistance into being packed within the separation

channel, even when being aided by the correct packing solution. The packing of both C18 bonded silica

and silica beads revealed to be impossible when also using this method, since although silica beads are

more hydrophilic, they are also smaller beads (58 µm average particle size and with a particle size

distribution of 40-75 µm) which will also reinforce the severely the increase in the pressure requirements

as seen in Equation 6, where b represents the particle size. To overcome this difficulty, a novel packing

method was employed: in order to pack the beads within the channel without the structure’s rupture or

leaking, the packing pressure had to be controlled and maintained within levels the microfluidic structure

could handle (between 0.15 and 0.2 MPa). This was achieved by pushing the beads suspension into

the channel by applying a constant pressure of 0.15 MPa using a compressed air line. Packing was

achieved after 30 minutes with two consecutive insertions of beads slurry since maintaining the packing

pressure leads to an eventual decrease in the flow velocity until packing stops and needs to be

reinitiated. Between insertions a PEG 20% solution is flowed to tightly pack the beads inside the channel.

Additionally, packing the separation channel two hours after performing a plasma cleaner treatment

demonstrated to facilitate the insertion particularly of the C18 beads, since the PDMS walls are still less

hydrophobic then they would be without any plasma cleaner treatment done to the surface. This packing

was adopted for further experiments with silica and C18 bonded silica beads since it was the fastest and

most efficient one, as seen in Figure 21 (A-D) where we can observe a fully packed 12 cm in length

separation channel.

In Figure 21 (left) we can observe a timeframe of the initial 12 seconds of silica beads packing

into the separation channel, highlighting the beads trapping feature already mentioned previously

(Section 3.1). This design provides trapping efficiencies of nearly 100% (Figure 21, timeframe and D).

However, beads are often heterogeneous in size which results, on rare occasions, in the smaller beads

being able to pass through the gap at the interface of the two channels as seen in Figure 21 (left,

timeframe) and therefore the extent to which this occurs does not impair the performance of the packing

step, since most of the time beads are entirely trapped in the separation channel (Figure 21, D). This

design allows the reduction of air bubbles formed inside the channels and provides mechanical

robustness, therefore allowing the device to withstand higher pressures than, for example, cage-like

PDMS device50, renewable and pneumatically controlled pillar array51 or a double weir design52, which

also increase the complexity of fabrication. Finally, the separation channels after packing could be used

even several weeks after, if properly stored under a DI water bath at around 4 ºC, to prevent the bed

from drying and formation of air bubbles.

43

3.4. Food colouring separation (proof-of-concept)

In order to test the chromatographic separation channel in a simple and straightforward way, a

colour-based separation was performed using the microfluidic structure from iteration 2 (simple cross-

channel injector without valves). A mixture of methylene blue, which holds a positive charge, and yellow

food colouring (which contains a azorubine and acetate, negatively charged) was successfully

separated (Figure 22, i-iv) with a packed column of sieved CM-Sepharose beads. CM-Sepharose is a

weak cation exchanger meaning that it is itself negatively charged.

Filtered CM-Sepharose bead suspension was prepared and packed inside the separation

channel. Then, Milli-Q water was flowed during about 10 minutes at a flow rate of 15 µL/min in order to

wash the separation channel from the components of the beads packing solution (Figure 22-A). A green

solution resulting from the mixture of methylene blue and yellow food colouring in a proportion of 1:1

was inserted on the cross-shaped injector inlet by flowing 3 μL/min perpendicularly to the separation

channel (Figure 22-B). Afterwards, Milli-Q water was used as mobile phase, with a flow rate of 8 μL/min.

(Figure 22-C). In Figure 22-(i), the green mixture had just been loaded and elution with Milli-Q water

Figure 22. Separation of a green coloured mixture. (A-C) corresponds to an illustration of the sequential steps

previously done to insertion of the mobile phase, including beads packing (A), green mixture solution loading (B)

and elution with Milli-Q water as the mobile phase (C). The microscopic images (i-iv) show the progression of the

separation throughout 10 minutes. (i) Initial sample plug starts to be eluted with Milli-Q water as mobile phase. (ii)

Methylene blue starts to be retained in the column. (iii) Yellow plug travels through the column. (iv) Eluted

components leave the column in a yellow plug with traces of green.

44

took place on the following steps (ii, iii, iv). As expected, the methylene blue was being retained by the

stationary phase. However, yellow food colouring (which contains quinoline yellow, azorubine and

acetate) is not being eluted alone and some of the methylene blue travels with the plug (Figure 22-iii),

partially acquiring a green colour. This may be explained by the fact that interaction between the

positively charged methylene blue and negatively charged azorubine can be partially happening. Still, a

great portion of the methylene blue was retained, proving the functionality of the chip. In Figure 22-iv,

the eluted components are leaving the column after about 10 minutes from the initial moment of elution.

It is important to note that as seen in Figure 22-i, the plug formed with the cross-channel injector

was indeed too wide for further separations on a more HPLC-like approach, which is the goal of this

work. This justifies the integration of valves within the optimized injector, that lead to iteration 4, later

used for other experiments in section 3.6.

Two main strategies were approached to proceed to the regeneration step (Figure 23) of the

separation channel: the first one was to try to equilibrate the charges of the methylene blue with buffers

of phosphate and different NaCl concentrations. At low ionic strength, competition between the buffer

ions and methylene blue for charged groups on the ion exchanger is minimal and methylene blue binds

strongly. Increasing the ionic strength increases the competition and so reduces the interaction between

the carboxymethyl group and methylene blue, causing it to elute. However, it revealed ineffective

washing the separation channel with phosphate buffer 1 M or 2 M NaCl solution in order to elute the

methylene blue since a continuous washing during 20 minutes at a flow rate of 13 µL/min did not wash

out the methylene blue from the channel.

Figure 23. Schematics illustrating the two employed strategies for separation channel regeneration: (1) increasing

the ionic strength of the mobile phase or (2) protonation of the carboxymethyl group of the stationary phase.

45

Anion and cation exchangers are classified as strong or weak, depending on how much the

ionization state of the functional groups varies with pH. A strong ion exchanger has the same charge

density on its surface over a broad pH range, whereas the charge density of a weak ion exchanger

changes with pH. Therefore, another approach was to protonate the carboxymethyl group present on

the beads by changing the pH, since it is classified as a weak cation exchanger. The carboxymethyl

group carries negative charge only above pH 4.5 (pKa: 4.0). Therefore, by changing the pH of the mobile

phase to below the pKa of the functional group with a HCl 1M solution flowed during 20 minutes at a

flow rate of 13 µL/min, the separation channel regeneration was possible. In Figure 23, the caption of

the microfluidic structure confirms this by showing the clean separation channel and a blue droplet at

the outlet, marked with an arrow.

3.5. Toxins separation for food safety applications

The developed microfluidic device was tested for food safety applications, namely the

separation of OTA and AFB1 toxins. In order to study the feasibility of separating these two toxins using

the microfluidic structure developed throughout this work, many factors had to be investigated, from the

toxins concentration to be used, the elution profiles of the toxins with different mobile phases, to the

actual elution using the 12 cm length separation channel and cross-channel injector.

3.5.1. Toxins concentration optimization

For the purpose of this experiment, the goal was to assess the optimum concentration of toxins

to use in further experiments, there was no need to use the long separation channels used so far for

this kind of assessment. Microchannels previously developed on the group56 were used for a rather fast

analysis of the parameters under study and packing, loading and elution were done as reported on the

literature. In Figure 24, the plots describing the variance of the fluorescence signal from the toxins OTA

and AFB1, and corresponding microscopic acquisitions are shown. OTA and AFB1 at different

concentrations were flowed by pulling into the microchannels previously packed with C18 bonded silica

beads and interacted differently with the beads based on the polarity of each toxin. It can be observed

that for OTA, a significative fluorescence signal can only be seen from a concentration higher than 3

µg/mL and that at 10 µg/mL the fluorescence signal is reasonably good, not being saturated. However,

for AFB1 the fluorescence signal is already relatively higher at 3 µg/mL when compared to the value of

OTA at the same concentration. At 10 µg/mL, the signal seems to be already saturated for AFB1.

Therefore, a compromising concentration of 6 µg/mL of both OTA and AFB1 was chosen to proceed to

the next experiments done with this microchannels.

46

3.5.2. OTA and AFB1 elution studies in microchannels

Acetonitrile and methanol are widely used in HPLC separations, mainly when using the RPC

mode, which also uses C18 bonded silica beads as a stationary phase. Acetonitrile is a polar aprotic

organic solvent, used as a medium-polarity solvent that is miscible with water and a range of organic

solvents. In reverse-phase separations, acetonitrile is generally considered the strongest eluent, above

methanol. However, for each separation, conditions have to be studied since the elution strength of a

compound is only a guideline to what will actually happen during the separation. In order to understand

which conditions of the mobile phase would be good to separate the two toxins in a separation channel

with the C18 bonded silica beads, again the microchannels were used to perform the experiments.

Similarly, to the previous section, solutions of OTA and AFB1 at 6 µg/mL were loaded and adsorbed to

the beads. Acetonitrile and methanol solutions of 75, 50, and 25% (v/v) in Milli-Q water were tested with

respect to the ability of eluting the adsorbed toxins. Acetonitrile and methanol solutions were flowed

Figure 24. Plots comprising the fluorescence signal variance in function of the concentration of OTA (top) and AFB1

(bottom) toxins. The correspondent microscopic images for the different concentrations for each toxin are displayed

on the left of the graphics. Fluorescence images were acquired at the beginning of the microchannel with an

exposure time of 1 second and a gain of 1x.

47

inside the microchannel by pushing and images were acquired every 30 seconds to analyse the

fluorescence signal. The plots obtained for the elution profiles can be seen in Figure 25 for both OTA

and AFB1.

Analysing the OTA elution profiles, it is clear that acetonitrile was stronger than methanol, eluting

almost entirely the OTA adsorbed on the beads, with the fluorescence signal reaching the same or lower

values when compared to the initial fluorescence at t=0 on any solvent ratio. The same analysis can be

applied for AFB1 elution profiles, where acetonitrile revealed to be a stronger eluent than methanol with

exception for the solvent ratio 25:75, where acetonitrile and methanol had similar elution strength.

Overall, the difference between the elution strength of acetonitrile and methanol was more accentuated

on OTA than in AFB1. Curiously, when observing the OTA elution plots for both acetonitrile and

methanol it is noticeable that the fluorescence signal seems to increase relatively to the initial loading

fluorescence during the first 1-2 minutes. This may be explained by a “wave-like” behaviour when

starting elution, since OTA molecules adsorbed on the beads that clogged on the inlet could travel to

the beginning of the microchannel and increase the fluorescence signal momently as they pass. To

overcome this from happening, caution should be taken to only pack beads on the microchannel and

avoid bead packing at the inlet. The presented fluorescence values on the plots were normalized

Figure 25. Elution profiles for toxins OTA and AFB1 previously adsorbed in a C18 bonded silica packed microchannel.

Elutions were performed during 10 minutes and fluorescence images acquired every 30 seconds at the beginning of

the microchannel with an exposure time of 1 second and a gain of 1x.

48

considering the background noise from the beads fluorescence. Also, the solvent in which toxins are

solubilized can alter the fluorescence signal of the toxins, increasing or decreasing it. This effect was

also considered and the presented plots are normalized taking that into account, although there was not

a significant change on the fluorescence signal that would alter the analysis of the plots (see section B,

Appendix 2)

Finally, a significant difference between the elution profile for OTA and AFB1 was observed for

a solvent ratio of CH3CN:water (25:75 v/v), where AFB1 was eluted more gradually than OTA, which

tended to stay on the mobile phase and quickly leave the microchannel. Therefore, this condition

promised some separation between the two toxins and was chosen to separate the toxins on the

experiments of the next section.

3.5.3. OTA and AFB1 cross-channel injection and elution

In order to study the feasibility of AFB1 elution within a packed column with a cross-channel

injector (iteration 2) and also the separation of AFB1 and OTA within the same column, the fluorescence

Figure 26. Cross-channel injection and elution of the toxins AFB1 and OTA in reverse-phase separation channel.

(A) illustrates the mode of operation for loading and elution of the toxins using iteration 2 as a microfluidic structure.

(B) are fluorescence acquisitions in the beginning of the separation channel comprising the cross-channel injector,

of the AFB1 6 µg/mL elution with 100% CH3CN (v/v) during the first 2 minutes. (C) are fluorescence acquisitions in

the beginning of the separation channel, of the elution with CH3CN:water (25:75 v/v) for the separation of OTA and

AFB1, at a concentration of 6 µg/mL Fluorescence images were acquired at the beginning of the microchannel with

an exposure time of 1 second and a gain of 1x.

49

of the toxins was followed in the reverse-phase packed channel (Figure 26). Before moving to directly

experimenting the toxins separation onto a microfluidic structure with a separation channel and cross-

channel injector with incorporated valves (iteration 4), it was important to assess the behaviour of the

toxin’s elution profiles within a long separation channel like the one developed throughout this work

before adding to the experiments the complex mechanism of the cross-channel injector with

incorporated valves. Therefore, the design from iteration 2 was used for this experiment, similarly to the

proof-of-concept coloured separation done previously. In Figure 26-A, the mode of operation for the

loading and elution of the toxins is shown. A volume of 60 µL of 6 µg/mL of AFB1 solution (for the elution)

or 60 µL of 6 µg/mL of AFB1 + OTA solution (for the separation) was loaded by pulling in the channel

perpendicular to the separation channel as seen in Figure 26-A. Acetonitrile 100% v/v (for the elution)

and 25% v/v (for the separation) in Milli-Q water solutions were flowed inside the microchannel by

pushing at the inlet for approximately 10 minutes.

In Figure 26-B we can observe the elution of AFB1 within the first 2 minutes with 100%

acetonitrile as mobile phase. The goal of this first experiment was to assess the behaviour of the

fluorescent plug while travelling through the separation channel, and therefore the strongest possible

eluent was used in pure form to ensure elution would happen. B-(i) refers to the moment where flowing

of the mobile phase had just started while in B-(ii, iii) the movement of the plug can already be observed.

In B-(iv), around two minutes after elution started the plug starts to spread and the fluorescence signal

to gradually faint until it was almost not distinguishable with the inherent fluorescent of the beads. This

limitation made following the fluorescent plug along the separation channel a really difficult task,

although a large volume of AFB1 solution (60 µL) was loaded in order to increase the initial fluorescent

of the plug and tackle this limitation. The plug continued to spread through the separation channel and

reached the outlet after around 10 minutes (results not shown), although the fluorescence signal was

so faint that the difference in the fluorescence signal cannot be regarded as 100 % sure belonging to

the plug. Nevertheless, the separation between OTA and AFB1 was also tried on this microfluidic

structure as seen in Figure 26-C. The expected outcome from this experiment was to at least observe

the initial plug starting to divide into two different fluorescent fronts somewhere at the beginning of the

separation channel while flowing CH3CN:water (25:75 v/v) as a mobile phase, since from the results of

the previous experiment it would be unlikely to be able to monitor the elution of the toxins at the end of

the separation channel due to the faint fluorescence signal. The mobile phase was chosen considering

the results from the experiments of section 3.5.2, where AFB1 would be more retained in the stationary

phase than OTA, at these chromatographic conditions. However, as seen in Figure 26-C, the initial plug

containing the two toxins gradually loses its fluorescent signal, not being clear if there were two fronts

of fluorescence being eluted although more than two minutes had passed between the first and last

image.

These results indicate three main problems that need to be overcome: first, the chromatographic

conditions need to be further optimized in order for the toxins to be effectively separated, especially the

mobile phase composition and flow rate used. An interesting approach would be adding acetic acid to

50

the composition of the mobile phase as demonstrated in the literature60. Secondly, the experimental

design to perform these experiments needs to be improved, since many limitations were experienced in

following the plug travelling through the separation channel and assess the actual difference between

the fluorescent signal of the toxins and the beads. Lastly, and most importantly, there is a clear problem

with both the design and packing of the separation channel causing band broadening as seen from the

dispersion the plug suffers while travelling through the packed channel. Recalling the van Deemter

equation (Equation 3):

𝐻 = 𝐶𝑀𝑢 (𝑜𝑟 𝐴) + 𝐵

𝑢+ 𝐶𝑆𝑢

One can speculate on which terms are probably affecting more the band broadening of the

separation channel given some known characteristics of the separation channel. For example, the term

A (eddy diffusion) refers to the mobile phase mass-transfer. Zone broadening in the mobile phase is

caused by the multiple paths a molecule can take through a packed column, especially when the lengths

of these paths can differ significantly. At low mobile phase velocities usually used in microfluidics, the

molecules are not significantly dispersed due to the contribution of regular diffusion effects that

attenuates the eddy diffusion. However, these multiple paths arise due to inhomogeneities in column

packing and small variations in the particle size of the packing material which is something to improve

in our separation channel, as discussed in section 3.1. For future work, is advised to use smaller particles

and with narrower size distribution that will lead to a better bed uniformity, to reduce the effect of this

parameter on band broadening. The term (𝐵𝑢⁄ ) represents longitudinal diffusion. This factor causes

band width to rise with time, and it happens whether or not the mobile phase is flowing. The time spent

by the band during its way through the column differs inversely with the flow rate, so the influence of

longitudinal diffusion on band width increases for low flow rates. Although in liquid chromatography,

diffusion rates are much smaller in value, so this term has an almost insignificant influence on band

broadening, this correlation may not be so straightforward in microfluidics as discussed in further section

3.6. Finally, the term (𝐶𝑆𝑢) represents the stationary phase mass-transfer: some analyte molecules will

enter further into a particle pore than others, spending in it a given and varied time. Molecules that spend

less time in the particle will move further along the column, with a resulting increase in band width. By

decreasing the particle size because of term A, the impact of this term will also be reduced, although

this term is probably the one least contributing to band broadening since dead volume is also a

contribution included in this effect and our microfluidic device was designed to be low dead volume

device.

Optimizing the chromatographic conditions and have a better experimental design are problems

that can be later addressed, independently of the design and packing of the separation channel.

However, studying the plug progression in order to assess the most important improvements to be done

in the future is imperative before tackling more advanced problems. Having this in mind, further

51

experiments were done in the next sections referring to the plug progression through the separation

channel.

3.6. Plug progression assessment and optimization

In order to study plug formation and progression inside a microfluidic structure with a cross-

shaped injector actuated by pneumatic valves (iteration 4B), plug optimization assays were performed

using FITC, a green fluorescent dye.

Figure 27 illustrates the necessary sequential steps in order to inject the plug and start elution.

In step 1 (washing), an ethanol 96% (v/v) solution was flowed by pushing using syringe pumps at a flow

rate of 15 µL/min for 20 minutes, from both the injector and separation channel inlets as indicated by

the black and white arrows, respectively. During this step, the valves from the loading channel of the

injector should be closed at both the inlet and outlet, as indicated by the black lines. This step is essential

to get rid on any air bubbles that may be entrapped between the injector and the separation channel

and, of course, remove any unwanted components from the channels. Next, the loading of 5 µL the

FITC 1mg/mL solution proceeded as seen in step 2 (loading) by pulling at 15 µL/min and subsequent

actuation of the loading channel valves while the other two were already closed (black lines). The plug

had a defined size, corresponding to the length between the two valves that give access to the

Figure 27. Schematics showing the operation of the cross-channel injector with integrated valves for the injection

and elution of the FITC plug by illustrating the sequential steps: washing (step 1), loading (step 2), injection (step

3) and elution (step 4). Arrows indicate the direction of fluid flow and dark lines the valves that are closed.

52

separation channel. Then, for the injection of the defined plug into the separation channel (step 3), the

valves that were previously closed were opened. Flow of the mobile phase should not start before

opening these valves that give access to the separation channel, since it was observed that pressure is

accumulated and when the valves are open, the accumulated pressure injects the plug too fast. Finally,

step 4 took place: keeping only the valves from the loading channel closed, the mobile phase (ethanol

96%) was flowed by pushing at a rate of 0.5 µL/min from both the injector and separation channel inlets,

illustrated by the white and black arrows, respectively. To note that in order for the elution to start, flow

had to be driven both from the injector and the separation channel inlets because flowing only from the

inlet, at the given flow rate, was not enough to win the accumulated fluid resistance in that zone of

intersection between the connection and separation channel. Therefore, one of the improvements in

future work is that the injector should not be placed perpendicular to separation channel in order to

ensure appropriate flow.

In Figure 28, a set of microscopic images are shown, that correspond to the sequential steps

described in Figure 27. From image (i) to (iii), the formation and injection of FITC plug can be observed.

Then, elution started (iv) in the separation channel without beads with ethanol 96% as the mobile phase

at a flow rate of 0.5 µL/min: the formation of a parabolic flow profile can immediately be seen. This

phenomenon is typical of pressure-driven separations and is regarded as the Taylor dispersion, where

a shear flow can increase the effective diffusivity of a given molecule. Shear acts to disperse the

concentration distribution in the direction of the flow, increasing the rate at which it spreads. The overall

effect is an enhancement of axial dispersion over molecular diffusion alone. From (v) to (vi) the evolution

of the plug observed corresponded to the expected evolution of plug under the effect of Taylor

dispersion. In (v) the spreading behaviour shortly after its injection time can be observed where the

cross-channel diffusion is yet to happen and the distance between the fastest moving molecules at the

centre and the slow particles on the wall stretches linearly with time. After some time (vi), enough to let

cross-channel diffusion to act, the edges of the paraboloid change to form a blob which is nearly uniform

Figure 28. Sequential fluorescent image acquisitions of a FITC plug injection (i-iii) and progression (iv-viii)

throughout the separation channel without beads packed. Images were acquired with an exposure time of 800

milliseconds and 3x gain.

53

across the channel and the plug spreads. In (vii) it can be seen the extension of the plug length after

traveling more than half of the separation channel, from the gradually decreased fluorescence signal in

each turn, which indicates that the plug can be several centimetres in length. Finally, in (viii) we can see

the front of the plug arriving at the outlet of the separation channel, after approximately 11 minutes had

passed since the beginning of the elution. From this experiment we could confirm the effect of Taylor

dispersion in our microfluidic device without beads packed within the separation channel. Additionally,

from image (v) to (vi) was a critical moment for the plug to disperse even more. This can be a

confirmation of the hypothesis reported in the literature, that factors such turns35,36 and corners37 can

influence plug spreading.

The question arises if this behaviour can also be observed when the separation channel is

packed with beads. Therefore, the same experiment was performed within a separation channel packed

with silica beads (Figure 29).

In Figure 29, a set of microscopic images are shown, that correspond to the sequential steps

described in Figure 27. From image (i) to (iii), the formation and injection of FITC plug can be observed.

Then, elution started (iv) in the separation channel with silica beads with ethanol 96% as the mobile

phase at a flow rate of 0.5 µL/min: the formation of a parabolic flow profile even in a packed channel

can be seen, although not as accentuated as in Figure 28-(iv). From (iv) to (v) the evolution of the plug

observed reached the “blob” state earlier in the separation channel, probably because of the beads

present within the channel. In (vi) and (vii), one can observe that similarly to the results without beads,

the spreading of the plug seems to be on the order of the several centimetres by analysing the different

intensities of fluorescence in each turn of the channel. Finally, in (viii) the front of the FITC plug reaches

the outlet around 13 minutes after elution started. This delay when compared to the same elution without

beads may be attributed to the beads itself, although FITC will interact shortly since it prefers more

Figure 29. Sequential fluorescent image acquisitions of a FITC plug injection (i-iii) and progression (iv-viii) throughout

the separation channel with beads packed. Images were acquired with an exposure time of 800 milliseconds and 3x

gain.

54

hydrophobic phases than silica. Although in liquid chromatography, diffusion rates are much smaller in

value, so this term has an almost insignificant influence on band broadening, this correlation may not

be so straightforward as demonstrated in this experiment. If a Taylor dispersion effect occurs in the

presence of beads, even if only in the beginning of the elution, it can hinder the separation efficiency of

the separation channel. Therefore, each microfluidic separation channel has to be regarded individually

and not much generalizations can be done in assessing the role of this effect on separation efficiency

and on the plug behaviour.

55

4. Conclusion and Future Prospects

Analytical separations can occur at any scale: from a laboratory scale to the industrial scale.

However, there is an increased demand for smaller and smaller instrumentation for analytical

separations5, requiring high-throughput answers to its problems but more importantly, fast and on-spot

analyses of products of interest. This implies small sample and set-up size, low costs and easy disposal

of wastes. In this experimental work, the main goal was to develop a miniaturized chromatographic

analytical device for food safety applications, which not only meets the goal of fully integration like

attempted previously12, but also aims for a simpler fabrication process (and therefore more economically

viable), user-friendly operation and versatility for a broad range of analytes/applications.

The main outcome of this work was the development of a microfluidic device which comprises a

cross-channel injector with integrated pneumatic actuated valves that offers almost zero dead-volume

and a 12 cm-long separation channel enabling the packing of most commercial available

chromatographic beads. The separation channel was tested for a proof-of-concept separation that

allowed the separation of a mixture of methylene blue and yellow food colouring proving the great

potential for future purification and analytical applications. Moreover, the applicability of this device for

food safety applications was assessed using a reverse-phase chromatographic mode for the separation

channel, indicating that under the optimal chromatographic conditions and further optimization of the

device design, separation and detection of toxins like AFB1 and OTA is possible.

Throughout the experimental work, five different microfluidic structures were developed from which

two represented the achievement of two of the main milestones of this project: first, iteration 2 for

establishing the 12 cm-long separation channel with an efficient beads trapping feature and lastly,

iteration 4B for providing an optimized cross-channel injector with integrated valves coupled to the

separation channel in a microfluidic structure that provided robustness for operation under relatively

high pressures for a soft-polymer based microfluidic device. Furthermore, optimizations regarding the

fabrication process were done that lead to optimized pneumatic actuation of the valves and the

application of a robust oxygen plasma cleaner interlayer bonding for multilayer soft lithography

fabricated devices. Nevertheless, although separation channel packing was achieved with different

commercially available chromatographic beads using a controlled-pressure packing method if smaller

beads or longer length for the separation channel might be needed, the microfluidic structure needs new

fabrication methodologies providing robustness to withstand the new pressure requirements. The beads

trapping feature adapted from previous work from the group56 revealed great performance in trapping

the beads of nearly 100%, even under a pressure accumulation from a 12-cm long packed channel

situation. However, again if smaller beads are aimed to be used this feature might need some adaption

to efficiently trap these small particles too. Regarding separations within the separation channel, a

mixture of methylene blue, which holds a positive charge, and yellow food colouring (which contains a

azorubine and acetate, negatively charged) was successfully separated with a packed column of sieved

56

CM-Sepharose beads (weak cation exchanger) with regeneration of the column possible by flowing HCl

1 M solution inside the channel. This proved the potential of the separation channel to provide both

preparative and analytical separation in several different applications, although some need for

improvements in the plug broadening right after insertion was observed because a valve-less cross-

channel injector was used. Nevertheless, the same separation channel was also tested for food safety

applications using a reverse-phase chromatographic mode for the separation channel, more precisely,

by eluting and separating OTA and AFB1. For this, separated elution studies for each toxin in

microchannels were performed to choose a mobile phase able to provide separation of the toxins. The

results from these experiments indicated that a deeper study of the chromatographic conditions needs

to be done and also further optimization of the device design since plug broadening was observed to

hinder the efficiency of separation. Regarding the plug broadening limitation observed, it was

hypothesized that the main players on this effect may be the mobile phase mass-transfer and

longitudinal diffusion phenomena present in the van Deemter equation. However, from studying the

FITC plug injection and progression in a separation channel with and without silica packing, it was

hypothesized that these limitations can be easily attained by providing a well-packed, smaller particle

packing and a more careful design as for example, a straight separation channel without turns or curves

in to dimming the effect of Taylor-dispersion.

Therefore, for future work, further optimization of the microfluidic device will be able to provide

better separations for reversed-phase chromatographic applications. More precisely, integration of

smaller particles (in the order of 5 µm), development of straight separation channel and consequently,

a fabrication process that delivers a device that can attain higher operation pressures while maintaining

the fabrication process simple and economically viable. For full integration, it will also be interesting to

integrate at the end of the separation channel a sensor in order to achieve the portability and fast

analysis aimed in LoC devices.

58

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Appendix Section

A. Optimized processes runsheet

Appendix 1. Runsheet for the final optimized process of the fabrication of iteration 4B, which includes the fluidic

layer and valves layer mold fabrication and also the whole optimized process for the PDMS structure fabrication.

STEP DESCRIPTION

SILICON SUBSTRATE

CLEANING

1. Silicon substrate 5x5 cm must be rinsed with acetone, followed

by IPA and finally DI water

2. Cleaned silicon substrate must be placed on a Petri dish filled

with Alconox solution, enough to fully cover the substrate

3. Put the Petri dish on bath at 65 ºC during 15 minutes

4. Clean the silicon substrate with DI water, followed by drying with

compressed air

5. Place the substrate on the UVO cleaner for 20 minutes

FLUIDIC LAYER MOLD

FABRICATION

6. SU-8 2015 photoresist spin-coating on top of the substrate

STEP 1: 10 seconds, 500 rpm, 100 rpm/s


7. Pre-exposure bake in hot plate at 95 ºC during 4 minutes with 1

minute for cooling down

8. Place the 20 µm design hard mask on top of the recently

deposited photoresist with the aluminium surface facing down

9. Exposure step to UV light during 30 seconds on the further way

slot

10. Post-exposure bake in a hot plate at 95 ºC during 5 minutes

with 2 minutes for cooling down

11. Development of the pattern by immersing the substrate in a

PGMEA solution during 2 minutes with manual agitation

12. Cleaning with IPA, followed by drying with compressed air

13. SU-8 50 photoresist spin-coating on top of the substrate



14. Pre-exposure bake in a hot plate by sequential baking

STEP 1: 65 ºC during 10 minutes

63

STEP 2: ramping up to 95 ºC


STEP 4: cooling down for 1 minute



16. Exposure step to UV light during 70 seconds on the further

way slot

17. Post-exposure bake in a hot plate by sequential baking

STEP 1: 65 ºC for 1 minute


STEP 3: 95 ºC for 10 minutes

STEP 4: cooling down for 2 minutes



REPEAT STEP 12

20. AZ 40XT spin-coating on top of the substrate



21. Let the spin-coated layer settle for 30 minutes


STEP 1: Slowly ramping from 100 to 125 ºC





24. Exposure step to UV light for 3 minutes and 30 seconds on the

further away slot

25. Post exposure in a hot plate at 105 ºC for 2 minutes with 2

minutes for cooling down

26. Development of the pattern by immersing the substrate in a AZ

400 K developer 1:3 DI water solution during 10 minutes with

manual agitation

27. Cleaning with DI water, followed by drying with compressed air

REPEAT STEP 22

VALVES LAYER MOLD

FABRICATION

28. SU-8 50 spin-coating on top of the substrate


64






STEP 4: cooling down for 1 minute



31. Exposure step to UV light for 25 seconds on the nearest slot

32. Post-exposure bake in a hot plate by sequential baking

STEP 1: 65 ºC during 1 minute






REPEAT STEP 12

34. Hard bake step in a hot plate at 150 ºC during 15 minutes

35. Slowly cooling down until 50 ºC

PDMS PREPARATION

36. PDMS elastomer and curing agent mixed in a 10:1 ratio

37. PDMS elastomer and curing agent mixed in a 20:1 ratio

38. Degasification step for 30 minutes in a desicator

PDMS CASTING

39. Casting for the fluidic layer channels by pouring the PDMS

preparation of 10:1 on top of the fluidic layer mold inside a Petri

dish

40. Casting for the valves layer channels by PDMS 20:1

preparation spin-coating on top of the valves layer mold at 25

seconds, 400 rpm and 100 rpm/s

41. PDMS curing by baking both in the oven at 70 ºC during 90

minutes

PDMS LAYERS BONDING

42. PDMS fluidic layer is cut and peeled off from the fluidic layer

mold

43. Holes are punched using 18 Gauge needles for the inlets and

outlets

44. Oxygen plasma bonding by placing both the PDMS fluidic layer

and PDMS valves layer in a oxygen plasma cleaner during 60

65

seconds at MED intensity, and sealed against each other

immediately after the plasma treatment

PDMS STACK SEALING

45. Holes are punched using 18 Gauge needles for the inlets of

the valves channels

46. PDMS 10:1 preparation spin-coating on top of a 1 mm-thick

glass slide



47. Oxygen plasma bonding by placing both the PDMS stack and

the PDMS spin-coated glass in a oxygen plasma cleaner during 60

seconds at MED intensity, and sealed against each other

immediately after the plasma treatment

66

B. Solvent effect influence on fluorescence signal for OTA

and AFB1 elution in microchannels

Appendix 2. Elution profiles for toxins OTA and AFB1 and respective curves normalized for the solvent effect on the

fluorescence signal of the toxins, previously adsorbed in a C18 bonded silica packed microchannel. Elution was

performed during 10 minutes and fluorescence images acquired every 30 seconds at the beginning of the microchannel

with an exposure time of 1 second and a gain of 1x. The elution profiles taking into account the solvent effect were

calculated by dividing the fluorescence values for each curve by the ratio between the toxin fluorescence signal at a given

mobile phase concentration and the same fluorescence signal in water. The curves were also normalized for the

background noise originated from the beads fluorescence.

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