Innovating Microfluidic Technologies

Jul 19, 2016
By LCGC Editors

The future of biological and clinical research will depend on technological innovations and cross discipline co-operation as science seeks a deeper understanding of increasingly complex biological systems. The 2016 recipient of the AES Mid-Career Award, Amy Herr, and her team at the University of California Berkeley have explored these areas using a combination of chemical engineering, mechanical engineering, and electrical engineering with strong foundations in biology, material science, and analytical chemistry to innovate new microfluidic analytical technology. She recently spoke to LCGC about this work. Interview by Lewis Botcherby


Q: Microfluidic analytical technology can reach scales—time-scales, length-scales, and concentrations—that are not possible to reach with macroscale tools. What do you consider the most important areas of research where these capabilities of microfluidic tools are proving most valuable?

A: Precision measurements. Using the capabilities of microfluidic design, we can measure miniscule spatial features and timescales (fast events) while at the same time making numerous measurements in parallel. For us, right now, these aspects come to bear in single-cell measurements, specifically multiplexed and high-specificity measurements across thousands of cells in a single assay. We have taken advantage of the precision to measure binding-induced conformational changes at the molecular level, for numerous conditions in a single assay. The throughput and precision are complementary, but they can be difficult to simultaneously achieve.

Q: One of the areas to which you have applied microfluidic analysis methods is to the study of riboswitches. Can you briefly explain what riboswitches are, and why you chose to study them?

A: Primarily found inside bacteria, riboswitches are RNA sensors that change conformation when the RNA molecule binds to a specific small-molecule metabolite. The conformation change of the RNA modulates gene expression. Discovered just a decade ago, only a small number of riboswitches have been identified, thus propelling further efforts at both discovery and characterization. An excellent example of current riboswitch research would be in the search for antibiotics. To circumvent issues with the development of antibiotic resistance, researchers are working on a new generation of antibiotics that target riboswitches as a way to kill the bacteria. Given the diversity of these molecules and the pioneering nature of the field, it is a thrilling time to be aiding efforts to discover riboswitches. The analytical tools required to detect and characterize these conformational changes in species are desperately needed, and the challenge of designing and developing these is incredibly rewarding.

Q: In 2013 you developed a microfluidic polyacrylamide gel electrophoresis (PAGE) technique—a microfluidic mobility shift assay (μMSA)—for the quantitative characterization of riboswitch–ligand binding interactions (1). How did this method compare to other methods for analyzing riboswitch–ligand binding?

A: Working with our collaborators, we realized early on that the discovery of riboswitches—and the cognate binding partners—would require a level of throughput suitable for screening. This brought a few design factors into focus, including the sparing consumption of the putative riboswitch molecules. We were also seeking to measure both binding and the concomitant change in RNA conformation. We needed to measure the “form” of the molecule to establish “function.” Electrophoretic mobility shift assays (EMSAs) are a perfect option, except that conventional slab-gel formats are neither fast nor highly parallelized—two attributes that can yield striking throughput. For us, that’s where the “μ” came in, “μ” meaning miniaturization. Miniaturization conferred both fast assay speed and a highly parallelized form factor. We employed classical enclosed microfluidic channels to demonstrate the EMSA assay speed in our 2013 paper(1), then followed that up with the twist of an open sieving structure (no enclosed microchannels) to support much higher parallelization in a follow-on study in 2014 (2). This enabled our group to harness the throughput and precision of microfluidic electrophoretic mobility shift assays to screen and validate five new candidate riboswitches in 0.3% of the time needed with existing tools.

Q: Two of the main goals of your μMSA method were to provide better resolution of the conformational changes that occur during riboswitch–ligand binding, and to enable riboswitch analysis without modifying the riboswitch aptamer structure. How did the method enable you to achieve these goals?

A: A really intriguing, and in hindsight, obvious outcome of the parallelized, miniaturized approach is the striking quantitative capacity of the new systems. The precision control of transport afforded by microfluidic formats gives a quantitative measure of the mobility shift. While that sounds great, the implications are actually quite striking. Instead of “eye balling” a shift on a slab gel, the parallelized device performs so many concurrent assays that we can now readily use statistical tests to establish the significance of conformation changes. Underpinning this quantitative leap is the control of electrophoresis, but also the ready ability to run multiple replicates on the same device and among different devices.

Q: Another goal of your μMSA method was to enable accurate measurement of both slow and rapid binding interactions (interconversion between bound and unbound forms). Were any modifications of the method necessary to obtain that information?

A: That’s right! The fast electrophoresis transport times made possible by miniaturization makes assessment of both rapidly and slowly interconverting riboswitch-metabolite pairs possible on the same device. Numerical simulations of the binding and electrophoresis transport timescales showed us that both long-lived complexes and more fleeting complexes are detectable with the rapid assays. In this way, the microfluidic format is relevant to the study of a much wider range of riboswitch binding pairs than conventional, macroscale assays. Really the modification was to move to rapid electrophoresis via miniaturization.

Q: In subsequent work, you developed an electrophoretic mobility shift assay (EMSA) suitable for high-throughput riboswitch screening (2). The research built on previous work that reported a 96-plex free-standing PAG electrophoresis (fsPAGE) assay (3). What challenges did you have to overcome to make the method work for high-throughput screening?

A: To get to the screening format, we had to compromise a little on miniaturization. Because we wanted numerous concurrent assays on one device, we moved away from classic “T-junction” channel networks as well as the sample and fluid reservoirs. In one way, we used the simple, straight discontinuous pore size gel to achieve low injection dispersion—and thus high performance separations—but in a single straight separation gel. We went with mesofluidic features, as they are substantially larger than capillary or microfluidic channel cross sections, because the largest gels can be integrated without complex fluidic networks and still support high electric field strength electrophoresis without Joule heating driving evaporation of the sample and run buffers before the separation completes.

Q: What were you able to achieve in terms of throughput?

Our published assays regarding validation of five new candidate riboswitches required just 0.3% of the time needed with existing tools. The performance translated into moving from 6000 seconds per data point with conventional tools to just 6 seconds per data point with the high throughput mesofluidic platform—a 1,000x reduction. Furthermore, sample consumption fell from 500 ng of RNA riboswitch material per data point to just 5 ng per data point, or a 100x reduction. Microfluidic design really can benefit throughput, which is relevant in molecular conformation change screening.

Q: In more recent work, you have developed a photoactive hydrogel to achieve single-cell resolution Western blotting (4). What were the advances you introduced to achieve this?

A: Our major contributions centered on creating a hydrogel that was both photo-activated and microchannel volume filling, with the former being achieved by incorporating benzophenone methacrylamide monomers. Most previous work had considered photoactive polymers for protein capture, but as surface coatings. A 3D reactive volume is quite different from a 2D reactive surface, with the volume based design not only offering more binding sites per channel length, but also reducing concerns of boundary layer depletion near the reactive 2D surface.

Q: What are the next steps in your research?

A: Currently, we are actively working to span the scales of biology with both high-throughput measurement tools and sub-cellular resolution tools. An overarching theme is our interest in assay and tool design regarding selective measurement of biomolecules, with quantitative capacity for somewhat nuanced physicochemical properties including modifications, complexes, conformation shifts, and activity. Intensive effort has been focused on single-cell resolution genomics and transcriptomics, however, focus has to be broader to also include single-cell resolution measurements of proteins, and their dynamic and critical forms and functions.

References

  1. K. Karns, J.M. Vogan, Q. Qin, S.F. Hickey, S.C. Wilson, M.C. Hammond, and A.E. Herr, J. Am. Chem. Soc. 135, 3136−3143 (2013).
  2. Y. Pan, T.A. Duncombe, C.A. Kellenberger, M.C. Hammond, and A.E. Herr, Anal. Chem. 86, 10357−10364 (2014).
  3. T.A. Duncombe and A.E. Herr, Lab Chip 13, 2115–2123 (2013).
  4. T.A. Duncombe, C-C. Kang, S. Maity, N Murthy, T.M. Ward, M.D. Pegram, and A.E. Herr, Adv. Mat. 28(2), 327-334 (2016).

Amy E. Herr received a BS degree in Engineering & Applied Science from the California Institute of Technology and MS and PhD degrees from Stanford University in Mechanical Engineering. She is currently the Lester John & Lynne Dewar Lloyd Distinguished Professor of Bioengineering at the University of California, Berkeley. Prior to joining UC Berkeley, she was a staff member in the Biosystems Research Group at Sandia National Laboratories (Livermore, CA). Her research interests include bioinstrumentation innovation needed to advance quantitation in life sciences and clinical problems, in particular the study and application of electrokinetic phenomena in multi-stage, heterogeneous bioanalytical microsystems. Her pedagogical interests are in bioengineering design, including innovation and translation.

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