Name: a microfluidic chip for icpms sample introduction
Uploaded: 2015-03-05T14:00:00.000+00:00
Duration: 11 min 16 s
Description: Watch this Scientific Journal Video about A Microfluidic Chip for ICPMS Sample Introduction at JoVE.com

This work was supported by the European Research Council (ERC Starting Grant n&#956;LIPIDS, No. 203428) and ETH Zurich (project number: ETH-49 12-2). The authors of this manuscript would like to thank Bodo Hattendorf for help with the ICP-MS and F. Kurth for cell counting. The authors also would like to thank Christoph B&#228;rtschi and Roland M&#228;der for their support with building the mechanical setup. The clean room facility FIRST at ETH Zurich is acknowledged for support in microfabrication.

1. SU-8 Master Fabrication (Figure 2)
NOTE: Perform the fabrication of the SU-8 master molds in a clean room to prevent defects caused by dust particles. Two wafers are needed for the fabrication, one wafer with microfluidic features and one without.
<ol>
	<li>Prepare the master molds for the microfluidic chip. First apply an adhesion layer to the silicon wafer.
	<ol>
		<li>Dehydrate a silicon wafer for 10&#160;min at 200&#160;&#176;C. Cool the wafer down to RT and load it on to a spin coater and spin coat it with SU-8 2002 with the following protocol.</li>
		<li>Dispense about 3&#160;ml resist onto the wafer.</li>
		<li>Spin the wafer at 500&#160;rpm for 10&#160;sec to spread the resist over the whole wafer.</li>
		<li>Spin the wafer at 2,000&#160;rpm for 30&#160;sec to achieve a resist height of approximately 2 &#181;m.</li>
	</ol>
	</li>
	<li>Remove excess resist from the edge of the wafer with an acetone soaked swab, to prevent sticking of the wafer to the hot plate in the next step. Bake the wafer for 60&#160;sec at 95&#160;&#176;C on a hot plate.</li>
	<li>Expose the whole wafer with ultraviolet light (80&#160;mJ/cm2 at 365&#160;nm). Post-bake the wafer for 120&#160;sec to 95&#160;&#176;C.</li>
	<li>Cool the wafer down and immediately spin coat the wafer again using the following protocol for SU-8 2050:
	<ol>
		<li>Spin the wafer at 100&#160;rpm for 20&#160;sec (dispense about 3&#160;ml SU-8 resist during this step).</li>
		<li>Spin the wafer at 500&#160;rpm for 10&#160;sec to spread the resist over the whole wafer.</li>
		<li>Spin the wafer at 3,250&#160;rpm for 30&#160;sec resulting in a resist thickness of approximately 40&#160;&#181;m.</li>
	</ol>
	</li>
	<li>Again, remove excess resist from the edge of the wafer with an acetone soaked swab and soft bake the wafer on a hot plate for 180&#160;sec at 65&#160;&#176;C and for 360&#160;sec at 95&#160;&#176;C.</li>
	<li>Prepare the photomask by sticking it to a soda-lime glass. See Figure 3 for the mask design. Use a mask aligner to expose the resist with ultraviolet light (160&#160;mJ/cm2, measured at 365 nm) through the prepared mask. Bake the exposed wafer again on a hot plate for 60&#160;sec at 65&#160;&#176;C and for 360&#160;sec at 95&#160;&#176;C.</li>
	<li>After cooling down the wafer to RT, immerse it in a glass Petri dish filled with developer for 5 min to develop the resist. Gently agitate the petri dish to remove unexposed SU-8. Rinse the wafer with isopropanol and blow it dry with a nitrogen gun.</li>
	<li>Examine the wafer under a microscope. In case undeveloped resist remains on the features, develop the wafer again for a few minutes, as described in step 1.7.</li>
	<li>Remove any residual solvent by baking the wafers for 2&#160;hr at 200&#160;&#176;C. Check the height of the features with a step profiler. In case the measured height differs from the desired height begin with this protocol again and adapt the spin speed in step 1.1.4.</li>
	<li>To prevent sticking of the PDMS to the wafer silanize it by placing it in desiccator together with 50&#160;&#181;l of 1H,1H,2H,2H-perfluorodecyltrichlorosilane in a small porcelain dish. Reduce the pressure in the desiccator to 100&#160;mbar and incubate the wafer for 12&#160;hr.
	<ol>
		<li>For the blank PDMS parts silanize another silicon wafer using the method of step 1.10. To save time silanize both wafers at the same time in a single desiccator.</li>
	</ol>
	</li>
</ol>
2. LADE Chip Fabrication
NOTE: The LADE chip is made out of two PDMS pieces that are bonded together by adhesive bonding24. The first part contains the microfluidic features. The other part is flat and used to seal the channels. Bonded together, they form the round shape necessary to interface the chip with the droplet transport system. Here, the fabrication of the two parts and their bonding is described. All process steps are shown in Figure 4.
<ol>
	<li>Prepare 44&#160;g of PDMS by mixing 40&#160;g of prepolymer with 4&#160;g of the PDMS curing agent (this will result in up to 6 chips). Degas the PDMS in a desiccator until it is bubble free (this will take about 20&#160;min).</li>
	<li>Replica molding of the structured halves.
	<ol>
		<li>Place the casting form on top of the wafer and snap it into place using the guiding structures around the design (see Figure 5). Skip the snapping into place for the flat PDMS halve.</li>
		<li>Pour approximately 3 to 4&#160;g of the degased PDMS in the casting form and place it for 6&#160;min on a hot plate at 150&#160;&#176;C. Cool down the cured PDMS in the casting form and carefully lift the casting form wafer using a spatula.</li>
		<li>In order to prevent any contamination of the microfluidic channels cover the side of the chip that was previously in contact with the wafer with tape. Carefully cut the tape along the edge of the PDMS part to remove excess PDMS.</li>
	</ol>
	</li>
	<li>To fabricate the flat PDMS halves repeat the above-mentioned steps 2.2.1 to 2.2.3 with the blank silanized wafer.</li>
	<li>Peel of the tape and punch fluid connection holes into the structured halves with a biopsy puncher. Protect the structures with tape during storage.</li>
	<li>Bond the PDMS parts together by adhesive bonding using the PDMS curing agent24.
	<ol>
		<li>Take an untreated silicon wafer and spin coat it with PDMS curing agent for 30&#160;sec at 6,000&#160;rpm. Take the wafer out of the spin coater.</li>
		<li>Remove the tape from the structured halves and place them with the structures facing downwards onto the wafer. Gently push on top of the PDMS to remove air bubbles.</li>
		<li>Remove the tape from the blank PDMS halves. Take a structured halve from the wafer and manually align it on top of the flat PDMS halve. Gently squeeze the part together to remove air bubbles and let the assembled chip cure for 24&#160;hr at RT. Do not push the parts together with force as this can cause the channels to collapse.</li>
	</ol>
	</li>
	<li>Cut the tip of the chip along the indicator line orthogonal to the nozzle channel with a utility knife to open the outlet nozzle. Use an alignment device to ensure a straight cut, which is necessary for a straight liquid ejection. Inspect the chip under a microscope for defects in the microfluidic channels and dust particles. Put a tape over the inlet holes to protect the chips during storage.</li>
	<li>Connect a Woulff bottle with tubing to a dry nitrogen source and to all inlets of the LADE chip. Deposit 50&#160;&#181;l of 1H,1H,2H,2H-perfluorodecyltrichlorosilane at the bottom of the Woulff bottle and close it.</li>
	<li>Silanize the microfluidic channels by flushing all channels for 20&#160;min with the nitrogen stream carrying 1H,1H,2H,2H-perfluorodecyltrichlorosilane at a flow rate of approximately 1&#160;ml/sec. The chips are ready for experiments and can be stored for at least several weeks at RT.</li>
</ol>
3. Preparations for Measurement / Droplet Transport System
NOTE: Build the whole droplet transport system on top of an optical table, since it is necessary to construct a stable supporting structure for the setup. A scheme of the whole droplet transportation system is depicted in Figure 6.
<ol>
	<li>Install a custom cyclonic poly(methyl methacrylate) (PMMA) adapter with an 50&#160;cm attached stainless steel tube vertically. Attach the adapter to a helium source with a mass flow controller. Attach a (high-speed) camera and a lamp to the adapter on the opposite sites for droplet visualization.</li>
	<li>Place a cartridge heater in the middle of the steel tube and use poly(vinyl chloride) (PVC) tubing and a Legris tube connector to connect the end of the steel tube with the inlet of the membrane desolvator.</li>
	<li>Connect the outlet of the desolvator with another PVC tubing to a laminar flow adapter, which is directly connected to the ICPMS inlet. Connect the laminar flow adapter to an argon source with a mass flow controller and later on use it to admix Argon to achieve a stable operation condition.</li>
	<li>Align the adapter as well as the steel tube vertical with a spirit level. If the alignment is not accurate, it may lead to significant losses of droplets. Insert a plug into the adapter to prevent gases leaking out during the system warm up time.</li>
	<li>Start all above-mentioned gas flows and devices using the settings from Table 1. Allow the system to warm up for 15 min. The cartridge heater needs 2 hr to stabilize the temperature, switch it on in advance.</li>
	<li>Place the syringe pumps on a rack at the height of cyclonic helium adapter. Keep the distance between the syringe pumps and the adapter as short as possible.</li>
</ol>
4. Measurements
NOTE: The following protocol is written in general terms because of the variety of solutions and suspensions that can be used. However, cell suspensions should be diluted to a concentration of &#60; 1 x 107 cells/ml, when single cell analysis is performed, to ensure that the majority of the droplets carry only one cell. For measurements with cells place the syringe pumps at an angle so that the outlet of the syringes point downwards and install the tubing in such a way that they point downwards.
<ol>
	<li>Attach tubing to the syringes. Load two 5&#160;ml syringes with perfluorohexane and one 1&#160;ml syringe with a sample solution or suspension. Remove all bubbles trapped in the syringes and tubing.</li>
	<li>Install the syringes in a syringe pump and connect them to the inlets of the chip. Start the syringe pumps using the start settings from Table 1 (or higher flow rates). Give the flows 3 to 5 min to stabilize.
	<ol>
		<li>Remove excess liquid from the tip of the chip with a tissue. The liquids should now eject from the chip in a straight jet. If a straight ejection cannot be achieved by wiping with a tissue replace the chip and start over with this step.&#160;</li>
	</ol>
	</li>
	<li>Remove the plug from the adapter and carefully insert the chip into the adapter. Lubricate the chip with FC-40 if necessary. A chip can be used for at least 2 hr of experiments.</li>
	<li>Change the flow rate to be within the recommended measurement settings from Table 1. Lower flow rate of the PFH not only saves PFH but also reduces signal background, caused by isobaric interferences.</li>
	<li>Give the system 2-5&#160;min to stabilize (depending on the chosen flow rates). Optimize the ICPMS for the highest signal intensity of the analytes of interest.
	<ol>
		<li>Successively adjust the flows of all the gases on the mass flow controllers (see the recommended ranges in Table 1) until the maximum signal intensity of the analytes of interest is achieved. Tune the plasma power and the focusing lens voltages (according to the manufacturer&#39;s recommended ranges) on the ICPMS in the same way.</li>
	</ol>
	</li>
	<li>Set the ICPMS to a dwell time of 10&#160;msec (applied for the very ICPMS used but can be adjusted with other instruments to ensure a time-resolved acquisition). Start recording the signal of a particular m/z using manufacturer&#8217;s protocol.</li>
	<li>After the measurement, transfer the raw data to data analysis program for evaluation. Bin the data, given in counts per 10 msec, with a built-in-function, and plot resulting bin center values against counts. Fit each peak in the plotted frequency distribution histogram with a Gauss function. The mean and sigma of the fit represent the mean signal intensity and its standard deviation, respectively.</li>
</ol>
5. Calibration Concept
<ol>
	<li>Measure a single or multi-element standard solution containing the element or elements of interest at the same flow rates as the sample.</li>
	<li>Place a LADE chip in a petri dish on a microscope. For a better image quality use a non-round chip. Fabricate this chip as described in step 2, but using a simple rectangular casting form instead of the partially round shaped one.</li>
	<li>Follow the steps 4.1 to 4.2.1 to start the droplet generation. Set the flow rates to the flow rates used in step 5.1.</li>
	<li>Record images of the aqueous droplets with a high-speed camera attached to a microscope (20X objective). Use an image analysis software like the droplet morphometry and velocimetry software by Basu25 to obtain the average droplet diameter from the recordings.</li>
	<li>Use average droplet diameter to calculate the droplet volume, assuming the droplet is a spherical object.
	<ol>
		<li>From this volume and the known concentration of an analyte in the droplet calculate the number of corresponding atoms. Divide the number of measured ions per droplet by the number of atoms to obtain the detection efficiency. Use this detection efficiency to calculate the number of atoms in an unknown sample. 
		NOTE: Since the variations between individual chips are small22, it is not necessary to repeat the measurement of the droplet size for every chip or solution if the flow rates remain the same. A list of the droplet sizes and frequencies according to the specific flow rates is published by Verboket et al.22.</li>
	</ol>
	</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pillar-induced+droplet+merging+in+microfluidic+circuits.">Pillar-induced droplet merging in microfluidic circuits.</a> Lab Chip. 8 (11), 1837-1841 (2008).</li><li>Song, H., Ismagilov, R. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Millisecond+kinetics+on+a+microfluidic+chip+using+nanoliters+of+reagents.">Millisecond kinetics on a microfluidic chip using nanoliters of reagents.</a> J. Am. Chem. Soc. 125 (47), 14613-14619 (2003).</li><li>Song, H., Chen, D. L., Ismagilov, R. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reactions+in+droplets+in+microflulidic+channels.">Reactions in droplets in microflulidic channels.</a> Angew. Chem., Int. Ed. 45 (44), 7336-7356 (2006).</li><li>Lombardi, D., Dittrich, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Droplet+microfluidics+with+magnetic+beads:+a+new+tool+to+investigate+drug-protein+interactions.">Droplet microfluidics with magnetic beads: a new tool to investigate drug-protein interactions.</a> Anal. Bioanal. Chem. 399 (1), 347-352 (2011).</li><li>Edgar, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compartmentalization+of+chemically+separated+components+into+droplets.">Compartmentalization of chemically separated components into droplets.</a> Angew. Chem., Int. Ed. 48 (15), 2719-2722 (2009).</li><li>Edgar, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capillary+electrophoresis+separation+in+the+presence+of+an+immiscible+boundary+for+droplet+analysis.">Capillary electrophoresis separation in the presence of an immiscible boundary for droplet analysis.</a> Anal. Chem. 78 (19), 6948-6954 (2006).</li><li>Baret, J. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence-activated+droplet+sorting+(FADS):+efficient+microfluidic+cell+sorting+based+on+enzymatic+activity.">Fluorescence-activated droplet sorting (FADS): efficient microfluidic cell sorting based on enzymatic activity.</a> Lab Chip. 9 (13), 1850-1858 (2009).</li><li>Agresti, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrahigh-throughput+screening+in+drop-based+microfluidics+for+directed+evolution.">Ultrahigh-throughput screening in drop-based microfluidics for directed evolution.</a> Proc. Natl. Acad. Sci. U. S. A. 107 (9), 4004-4009 (2010).</li></ol>

The authors declare that they have no competing financial interests.

Although the fabrication of the chips is very reliable there are some critical points during the fabrication that require special attention. First, cleanliness during the assembly is highly important to prevent contamination of the chip by dust. The dust can block the channels and prevent a stable droplet generation. Second, it is especially important that the tip is cut orthogonal to the nozzle channel. The angle of the cut strongly influences the ejection angle. If the liquid is ejected at an angle it can cause a loss of the ejected droplets.
When building the setup ensure that it is stable. The vertical alignment of the metal tube and the adapter are important. Also during the measurements there are some points that need special attention. The insertion of the chip into the adapter has to be performed carefully. It can happen that during the insertion the jet is disrupted and stops. For measurements with cells the position and orientation of the syringe pumps and tubing are important. Their proper placement can reduce settling of the cells in the syringe and the tubing.
The LADE chip presented here has several advantages over existing commercial droplet generators. The system is more robust, provides a wider droplet size range, which can be further extended by modification of the channel geometry, and is disposable. A single use device is of a particular interest for the analysis of samples with a high content of salts or solid residues, as for instance nanoparticles or cell suspensions, which can clog tiny channels and cannot always be easily washed out. The transport of single microdroplets generated by the LADE into the MS is still a limiting step in our system and has to be further optimized. The current droplet transport assembly, although removes the PFH vapor, which would otherwise create additional spectral and non-spectral interferences and cause plasma instabilities, but still results in a high temporal jitter of droplet arrival at the MS and an incomplete droplet transport. In comparison to the commercial available droplet introduction systems the transport system for this setup requires more equipment. The current chip is designed for sample introduction only. However, with slight changes of the design advanced introduction and sample preparation steps cloud be implemented on-chip, e.g., dilution27-29, ultra fast mixing30, chemical reactions31, separation32-34, or cell sorting35,36. Under advanced introduction devices we understand for example the introduction of sample and standard droplets sequentially or parallel with a single chip. This would increase the throughput and improve the accuracy of quantitative analysis.

Elemental analysis of liquid samples by inductively coupled plasma mass spectrometry (ICPMS) is commonly carried out using nebulizers in combination with spray chambers as introduction system1. In this sample introduction system the sample is sprayed by a nebulizer to generate a polydisperse aerosol. A downstream spray chamber is used to filter out large droplets. This method is associated with high sample consumption (&#62;0.3 ml min-1)2 and an incomplete sample transport. Thus, it becomes impractical for applications where only microliter sample volumes are available, as in biological, forensic, toxicological and clinical studies3. To reduce the sample consumption, nebulizers with smaller nozzle dimensions were developed3. However, the reduced nozzle size increases the risk of clogging when samples of undigested biological fluids or concentrated salt solutions have to be analyzed3.
A different approach for sample introduction was proposed by Olesik et al.4. The authors injected a liquid into ICPMS in the form of monodisperse discrete microdroplets, which were produced by a piezo-electrically driven micropump. Even though this very system did not find wide application, it initiated the further development of the concept of discrete droplet introduction in ICPMS. Today, piezo-electrically driven dispensing systems, which can generate droplets in size of 30, 50, 70 and 100 &#181;m and at frequencies of 100-2,000 Hz, can be purchased. The droplets can be transported into ICPMS with close to 100% efficiency5. These microdroplet dispensers have been applied for quantitatively measuring single nanoparticles5,6 as well as characterizing individual biological cells7. A similar system based on thermal inkjet technology8 was tested for analysis of biological samples9. Although the available single droplet introduction systems are very efficient, can be used for small sample volumes and are promising for the analysis of nanoparticles and cells, they have several limitations. For a fixed nozzle size, the droplet size can be varied only slightly (unless custom settings are used10). Changes of the physical properties of the liquid (pH, salt content) can alter the droplet characteristics (size, injection speed). Also, these devices are rather expensive, prone to clogging and are difficult to clean.
Another method to generate droplets is known in the field of droplet microfluidics11. In recent years droplet microfluidics has gained interest for (bio-)chemical reactions12-15 and for single cell studies16,17. Additionally, this technique was applied for introducing samples in electrospray ionization mass spectrometry18,19 and for preparing samples in matrix-assisted laser desorption/ionization mass spectrometry20,21.
Recently, we introduced a microfluidic based system for sample introduction in ICPMS22. The key component of our introduction system is the liquid assisted droplet ejection (LADE) chip. This chip consists completely of poly(dimethylsiloxane) (PDMS). In the first channel junction flow focusing is used to generate monodisperse droplets of an aqueous sample solution (Figure 1). For this purpose the highly volatile (boiling point of 58-60&#160;&#176;C23) and immiscible carrier phase perfluorohexane (PFH) is used (Figure 1). These PFH properties enable a stable droplet generation and fast removal of the carrier phase. Changes in the properties of the sample liquid influence this generation method less, compared to other droplet generators. The droplet size is adjustable over a wide range by changing the flow rates of the aqueous phase and the PFH. In a downstream secondary junction, more PFH is added to increase the flow speed to at least 1 m&#160;sec-1. At this speed the liquid can be ejected from the chip in stable and straight jet (Figure 1) without droplet destruction (Figure 1 inset). This double-junction design allows controlling the jet stability independent of droplet generation. The droplets are transported to the ICPMS with a customized transport system. This system comprises a falling tube and a membrane desolvator to remove the PFH. The dried residues of the aqueous droplets are subsequently ionized in the plasma of the ICPMS and a mass detector measures the ions. The front part of the chip is barrel-shaped to ensure a tight connection with the droplet transport system. The ejection of the aqueous sample as droplets in PFH is beneficial, because contact with the nozzle is avoided. This considerably lowers the risk of nozzle clogging, which can be a problem when working with cell suspensions or concentrated salt solutions. The LADE chips, fabricated by PDMS soft lithography, are cheap (material cost approximately $2 per chip), disposable and easy to modify. In combination with the fabrication that requires only a small amount of manual work each experiment can be performed with a new chip. Therefore, a laborious cleaning is not needed and cross contamination is minimized.
Here, the fabrication of the LADE chip by soft lithography and its application for ICPMS are described. Examples of measurements with an aqueous solution and a cell suspension are presented.

<table><tbody><tr><td>Silicon wafer 100 mm</td><td>Si-Mat (Kaufering, Germany)</td><td></td><td></td></tr><tr><td>SU-8 2002</td><td>Microchem Corp. (Massachusetts, U.S.A.)</td><td></td><td></td></tr><tr><td>SU-8 2050</td><td>Microchem Corp. (Massachusetts, U.S.A.)</td><td></td><td></td></tr><tr><td>Acetone</td><td>Merk VWR (Darmstadt, Germany)</td><td>100014</td><td></td></tr><tr><td>MR-developer 600</td><td>Microresist Technology GmbH (Berlin, Germany)</td><td></td><td></td></tr><tr><td>Isopropanol</td><td>Merk VWR (Darmstadt, Germany)</td><td>109634</td><td></td></tr><tr><td>1&lt;em&gt;H&lt;/em&gt;,1&lt;em&gt;H&lt;/em&gt;,2&lt;em&gt;H&lt;/em&gt;,2&lt;em&gt;H&lt;/em&gt;-perfluorodecyltrichlorosilane</td><td>ABCR-Chemicals (Karlsruhe, Germany)</td><td>AB111155</td><td></td></tr><tr><td>Sylgard 184 silicone elastomer kit (PDMS)</td><td>Dow Corning (Michigan, U.S.A.)</td><td>39100000</td><td></td></tr><tr><td>Perfluorohexane 99%</td><td>Sigma-Aldrich (Missouri, U.S.A.)</td><td>281042</td><td></td></tr><tr><td>FC-40</td><td>ABCR-Chemicals (Karlsruhe, Germany)</td><td>AB103511</td><td></td></tr><tr><td>Phosphate-buffered saline&amp;nbsp;</td><td>Life Technologies (Paisley, U.K.)&amp;nbsp;</td><td>10010-015</td><td></td></tr><tr><td>Red blood cells in phosphate-buffered saline</td><td>Rockland Immunochemicals Inc. (Pennsylvania, U.S.A.)&amp;nbsp;</td><td>R400-0100</td><td></td></tr><tr><td>Single-element standard solutions Na, Fe</td><td>Inorganic Ventures (Virginia, U.S.A.)</td><td></td><td></td></tr><tr><td>Multielement standard solution&amp;nbsp;</td><td>Merck Millipore (Massachusetts, U.S.A.)</td><td>IV</td><td></td></tr><tr><td>Nitric acid</td><td>Sub-boiled</td><td></td><td></td></tr><tr><td>Ultrahigh-purity water</td><td>Merck Millipore (Massachusetts, U.S.A.)</td><td></td><td></td></tr><tr><td>Hot plate HP 160 III BM</td><td>Sawatec (Sax, Switzerland)</td><td></td><td>used for wafer preparation</td></tr><tr><td>Spin modules SM 180 BM</td><td>Sawatec (Sax, Switzerland)</td><td></td><td>used for wafer preparation</td></tr><tr><td>High resolution film photomask</td><td>Microlitho (Essex, U.K.)</td><td></td><td></td></tr><tr><td>Step profiler Dektak XT advanced</td><td>Bruker&amp;nbsp; (Massachusetts, U.S.A.)</td><td></td><td></td></tr><tr><td>Hot plate MR 3002</td><td>Heidolph (Schwabach, Germany)</td><td></td><td>used for replica molding&amp;nbsp;</td></tr><tr><td>1.5 mm biopsy puncher</td><td>Miltex (Pennsylvania, U.S.A.)</td><td>33-31AA/33-31A</td><td></td></tr><tr><td>Spin coater&amp;nbsp; WS-400 BZ-6NPP/LITE</td><td>Laurell (Pennsylvania, U.S.A.)</td><td></td><td>used for adhesive bonding</td></tr><tr><td>Syringe pump neMESYS</td><td>Cetoni (Korbussen, Germany)</td><td></td><td></td></tr><tr><td>1 ml syringe&amp;nbsp;</td><td>Codan (Lensahn, Germany)&amp;nbsp;</td><td>62.1002</td><td></td></tr><tr><td>5 ml syringe&amp;nbsp;</td><td>B. Braun (Melsungen, Germany)&amp;nbsp;</td><td>4606051V</td><td></td></tr><tr><td>PTFE tubing&amp;nbsp;</td><td>PKM SA (Lyss, Switzerland)&amp;nbsp;</td><td>PTFE-AWG-TFT20.N</td><td></td></tr><tr><td>Quadrupole-based ICPMS ELAN6000</td><td>PerkinElmer (Massachusetts, U.S.A.)&amp;nbsp;</td><td></td><td></td></tr><tr><td>Membrane desolvator CETAC6000AT+</td><td>CETAC Technologies (Nebraska, U.S.A.)&amp;nbsp;</td><td></td><td>only the desolvator unit is used</td></tr><tr><td>High speed camera Miro M110</td><td>Vision Research (New Jersey, U.S.A.)</td><td></td><td></td></tr><tr><td>Data analysis program Origin pro</td><td>OriginLab Corp. (Massachusetts, U.S.A.)</td><td>version 8.6</td><td></td></tr><tr><td>Microscope</td><td>Olympus (Tokyo, Japan)</td><td>IX71</td><td></td></tr></tbody></table>

a microfluidic chip for icpms sample introduction

This protocol discusses the fabrication and usage of a disposable low cost microfluidic chip as sample introduction system for inductively coupled plasma mass spectrometry (ICPMS). The chip produces monodisperse aqueous sample droplets in perfluorohexane (PFH). Size and frequency of the aqueous droplets can be varied in the range of 40 to 60 &#181;m and from 90 to 7,000&#160;Hz, respectively. The droplets are ejected from the chip with a second flow of PFH and remain intact during the ejection. A custom-built desolvation system removes the PFH and transports the droplets into the ICPMS. Here, very stable signals with a narrow intensity distribution can be measured, showing the monodispersity of the droplets. We show that the introduction system can be used to quantitatively determine iron in single bovine red blood cells. In the future, the capabilities of the introduction device can easily be extended by the integration of additional microfluidic modules.

The presented system can be employed to measure small volumes of solutions or suspensions containing cells or nanoparticles. Examples of a measurement of a standard solution and characterization of single cells are shown here. More examples can be found in Verboket et al.22.
Typically the signal of a single droplet of a solution is a very short event. It usually lasts for a few 100 &#181;sec26. With the ICPMS used here (dwell time 10 msec) short signals like these cannot be temporally resolved. Figure 7a and Figure 7b show the signals and frequency distribution histogram of a Na standard solution. The droplets arrive at the plasma with a temporal jitter &#62; 10 msec. The detection is unsynchronized. Signals of one (201 &#177; 24), two (381 &#177; 34) or three (560 &#177; 45) droplets are detected within one dwell time. Low variation in signal intensity suggests high droplet monodispersity. The first tailing peak is likely the result of droplet fragments; the cause of this fragmentation is still under investigation.
The calibration approach described in 5 (using Fe standard solution) was tested for the determination of Fe content of single bovine/calf red blood cells (6-7 &#181;m in diameter) suspended in phosphate buffered saline (PBS). The suspension of 1 x 107 cells ml-1 was used to ensure that the majority of droplets carry only one cell. Figure 8 shows the signals from cells as frequency distribution histogram. On average every cell contained 5.3 &#177; 1.2 x 108 Fe atoms (see Verboket et al.22).
<img alt="Figure 1" src="/files/ftp_upload/52525/52525fig1highres.jpg" /> 
Figure 1. Micrograph of droplet generation, acceleration and ejection. In a flow focusing junction, monodisperse aqueous droplets are generated in the stream of PFH. Additional PFH is added at a second junction. Subsequently, the liquid stream is ejected from the LADE chip through a nozzle. Arrows indicate the direction of the flow. Scale bar is 500 &#181;m. Inset: Micrograph of an aqueous droplet in a PFH shell after ejection from the LADE chip. Scale bar 100 &#181;m. Adapted with permission from22. Copyright 2014 American Chemical Society. This figure has been modified with data from research conducted since publication in the laboratory of P. S. Dittrich. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52525/52525fig2highres.jpg" /> 
Figure 2. Schematic of the SU-8 processing. First an adhesion layer is coated. A silicon wafer is spin coated with SU-8 2002, soft baked, flood exposed with ultraviolet light and post baked. On top of this layer the microfluidic structures are produced. The wafer is spin coated with SU-8 2050 and soft baked. The design of the microfluidic structures is transferred to the wafer by exposing it with ultraviolet light through a photomask. After a postexposure bake, the photoresist is developed and a hardbake is performed. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52525/52525fig3highres.jpg" /> 
Figure 3. Design of the photomask for the LADE chip containing the following features: a) guiding structures for the casting form, b) an inlet for the PFH for droplet acceleration, c) an inlet for PFH for droplet generation and d) an inlet for the aqueous sample. e) Indicator line for cutting off the tip of the chip. Channel widths f) = 40 &#181;m, g) = 20 &#181;m and h) = 25 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52525/52525fig4highres.jpg" /> 
Figure 4. Process chart of the LADE chip fabrication. First the structured and the flat PDMS parts are fabricated by replica molding. The two pieces are bonded together by adhesive bonding. Finally, the tip of the chip is cut off and the microfluidic channels are silanized. Adapted with permission from22. Copyright 2014 American Chemical Society. This figure has been modified since publication. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52525/52525fig5highres.jpg" /> 
Figure 5. Technical drawing of the aluminum casting form for the LADE chip. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/52525/52525fig6highres.jpg" /> 
Figure 6. Schematic drawing of the setup (not to scale). The system consists of the LADE chip, a cyclonic adapter, a heated steel tube, a membrane desolvator, and an ICPMS. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/52525/52525fig7highres.jpg" /> 
Figure 7. (A) Signals of droplets made from of a 1 mg kg-1 Na standard solution. (B) Frequency distribution histogram of these signals. In the 10 msec dwell time signals of one (yellow), two (red) or three (blue) droplets were recorded. Mean and standard deviation of the signals were determined by fitting Gaussian functions. The flow rates used were 0.5, 50, and 60 &#181;l min&#8722;1 of aqueous sample, PFH for droplet generation, and PFH for droplet acceleration, respectively. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52525/52525fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" src="/files/ftp_upload/52525/52525fig8highres.jpg" /> 
Figure 8. Frequency distribution histogram of 56Fe+ signals generated by red blood cells. Mean and standard deviation of the signals were determined by fitting a Gaussian function. The flow rates used were 2, 80, and 80 &#181;l min&#8722;1 of cell suspension, PFH for droplet generation, and PFH for droplet acceleration, respectively.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>He gas flow rate</td>
			<td></td>
			<td>0.6-0.8 L min-1</td>
		</tr>
		<tr>
			<td>Cartridge heater</td>
			<td></td>
			<td>30 W</td>
		</tr>
		<tr>
			<td>Desolvator membrane temperature</td>
			<td></td>
			<td>160 &#176;C</td>
		</tr>
		<tr>
			<td>Desolvator sweep gas flow rate</td>
			<td></td>
			<td>3-4 L min-1</td>
		</tr>
		<tr>
			<td>Ar gas flow rate</td>
			<td></td>
			<td>0.1 L min-1</td>
		</tr>
		<tr>
			<td>ICP plasma power</td>
			<td></td>
			<td>1,300 W</td>
		</tr>
		<tr>
			<td>Sample flow rate</td>
			<td>start up</td>
			<td>1 &#181;l min-1</td>
		</tr>
		<tr>
			<td></td>
			<td>measurement</td>
			<td>0.3-15 &#181;l min-1</td>
		</tr>
		<tr>
			<td>Flow rate of PFH droplet generation</td>
			<td>start up</td>
			<td>60 &#181;l min-1</td>
		</tr>
		<tr>
			<td></td>
			<td>measurement</td>
			<td>35-80 &#181;l min-1</td>
		</tr>
		<tr>
			<td>Flow rate of PFH droplet acceleration</td>
			<td>start up</td>
			<td>60 &#181;l min-1</td>
		</tr>
		<tr>
			<td></td>
			<td>measurement</td>
			<td>35-80 &#181;l min-1</td>
		</tr>
	</tbody>
</table>
Table 1. Start settings and recommended measurement settings for the ICPMS and the syringe pumps.

Watch this Scientific Journal Video about A Microfluidic Chip for ICPMS Sample Introduction at JoVE.com

A Microfluidic Chip for ICPMS Sample Introduction

We present a discrete droplet sample introduction system for inductively coupled plasma mass spectrometry (ICPMS). It is based on a cheap and disposable microfluidic chip that generates highly monodisperse droplets in a size range of 40&#8722;60 &#181;m at frequencies from 90 to 7,000 Hz.

This protocol discusses the fabrication and usage of a disposable low cost microfluidic chip as sample introduction system for inductively coupled plasma ...

a-microfluidic-chip-for-icpms-sample-introduction

Nanyang Technological University

Research

JoVE Journal

Bioengineering

11.1K Views.  ETH Zurich.  We presenteren een discrete druppel monster introductie voor inductief gekoppeld plasma massaspectrometrie (ICPMS). Het is gebaseerd op een goedkope en wegwerpbare microfluïdische chip die sterk monodisperse druppeltjes in een groottebereik van 40-60 pm bij frequenties van 90 tot 7000 Hz. 

Video: A Microfluidic Chip for ICPMS Sample Introduction

A Microfluidic-based Hydrodynamic Trap for Single Particles

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science. Methods for particle trapping based on optical, magnetic, electrokinetic, and acoustic techniques have led to major advancements in physics and biology ranging from the molecular to cellular level. In this article, we introduce a new microfluidic-based technique for particle trapping and manipulation based solely on hydrodynamic fluid flow. Using this method, we demonstrate trapping of micro- and nano-scale particles in aqueous solutions for long time scales. The hydrodynamic trap consists of an integrated microfluidic device with a cross-slot channel geometry where two opposing laminar streams converge, thereby generating a planar extensional flow with a fluid stagnation point (zero-velocity point). In this device, particles are confined at the trap center by active control of the flow field to maintain particle position at the fluid stagnation point. In this manner, particles are effectively trapped in free solution using a feedback control algorithm implemented with a custom-built LabVIEW code. The control algorithm consists of image acquisition for a particle in the microfluidic device, followed by particle tracking, determination of particle centroid position, and active adjustment of fluid flow by regulating the pressure applied to an on-chip pneumatic valve using a pressure regulator. In this way, the on-chip dynamic metering valve functions to regulate the relative flow rates in the outlet channels, thereby enabling fine-scale control of stagnation point position and particle trapping. The microfluidic-based hydrodynamic trap exhibits several advantages as a method for particle trapping. Hydrodynamic trapping is possible for any arbitrary particle without specific requirements on the physical or chemical properties of the trapped object. In addition, hydrodynamic trapping enables confinement of a "single" target object in concentrated or crowded particle suspensions, which is difficult using alternative force field-based trapping methods. The hydrodynamic trap is user-friendly, straightforward to implement and may be added to existing microfluidic devices to facilitate trapping and long-time analysis of particles. Overall, the hydrodynamic trap is a new platform for confinement, micromanipulation, and observation of particles without surface immobilization and eliminates the need for potentially perturbative optical, magnetic, and electric fields in the free-solution trapping of small particles.

In this article, we present a microfluidic-based method for particle confinement based on hydrodynamic flow. We demonstrate stable particle trapping at a fluid stagnation point using a feedback control mechanism, thereby enabling confinement and micromanipulation of arbitrary particles in an integrated microdevice.

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science.  Methods for ...

Microfluidic Chips Controlled with Elastomeric Microvalve Arrays

Miniaturized microfluidic systems provide simple and effective solutions for low-cost point-of-care diagnostics and high-throughput biomedical assays. Robust flow control and precise fluidic volumes are two critical requirements for these applications. We have developed microfluidic chips featuring elastomeric polydimethylsiloxane (PDMS) microvalve arrays that: 1) need no extra energy source to close the fluidic path, hence the loaded device is highly portable; and 2) allow for microfabricating deep (up to 1 mm) channels with vertical sidewalls and resulting in very precise features.

The PDMS microvalves-based devices consist of three layers: a fluidic layer containing fluidic paths and microchambers of various sizes, a control layer containing the microchannels necessary to actuate the fluidic path with microvalves, and a middle thin PDMS membrane that is bound to the control layer. Fluidic layer and control layers are made by replica molding of PDMS from SU-8 photoresist masters, and the thin PDMS membrane is made by spinning PDMS at specified heights. The control layer is bonded to the thin PDMS membrane after oxygen activation of both, and then assembled with the fluidic layer. The microvalves are closed at rest and can be opened by applying negative pressure (e.g., house vacuum). Microvalve closure and opening are automated via solenoid valves controlled by computer software.

Here, we demonstrate two microvalve-based microfluidic chips for two different applications. The first chip allows for storing and mixing precise sub-nanoliter volumes of aqueous solutions at various mixing ratios. The second chip allows for computer-controlled perfusion of microfluidic cell cultures.

The devices are easy to fabricate and simple to control. Due to the biocompatibility of PDMS, these microchips could have broad applications in miniaturized diagnostic assays as well as basic cell biology studies.

We demonstrate protocols for manufacturing and automating elastomeric polydimethylsiloxane (PDMS)-based microvalve arrays that need no extra energy to close and feature photolithographically defined precise volumes. A parallel subnanoliter-volume mixer and an integrated microfluidic perfusion system are presented.

Miniaturized microfluidic systems provide simple and effective solutions for low-cost point-of-care diagnostics and high-throughput biomedical assays.  ...

Biology

On-chip Isotachophoresis for Separation of Ions and Purification of Nucleic Acids

Electrokinetic techniques are a staple of microscale applications because of their unique ability to perform a variety of fluidic and electrophoretic processes in simple, compact systems with no moving parts. Isotachophoresis (ITP) is a simple and very robust electrokinetic technique that can achieve million-fold preconcentration1,2 and efficient separation and extraction based on ionic mobility.3 For example, we have demonstrated the application of ITP to separation and sensitive detection of unlabeled ionic molecules (e.g. toxins, DNA, rRNA, miRNA) with little or no sample preparation4-8 and to extraction and purification of nucleic acids from complex matrices including cell culture, urine, and blood.9-12
ITP achieves focusing and separation using an applied electric field and two buffers within a fluidic channel system. For anionic analytes, the leading electrolyte (LE) buffer is chosen such that its anions have higher effective electrophoretic mobility than the anions of the trailing electrolyte (TE) buffer (Effective mobility describes the observable drift velocity of an ion and takes into account the ionization state of the ion, as described in detail by Persat et al.13). After establishing an interface between the TE and LE, an electric field is applied such that LE ions move away from the region occupied by TE ions. Sample ions of intermediate effective mobility race ahead of TE ions but cannot overtake LE ions, and so they focus at the LE-TE interface (hereafter called the "ITP interface"). Further, the TE and LE form regions of respectively low and high conductivity, which establish a steep electric field gradient at the ITP interface. This field gradient preconcentrates sample species as they focus. Proper choice of TE and LE results in focusing and purification of target species from other non-focused species and, eventually, separation and segregation of sample species.
We here review the physical principles underlying ITP and discuss two standard modes of operation: "peak" and "plateau" modes. In peak mode, relatively dilute sample ions focus together within overlapping narrow peaks at the ITP interface. In plateau mode, more abundant sample ions reach a steady-state concentration and segregate into adjoining plateau-like zones ordered by their effective mobility. Peak and plateau modes arise out of the same underlying physics, but represent distinct regimes differentiated by the initial analyte concentration and/or the amount of time allotted for sample accumulation.
We first describe in detail a model peak mode experiment and then demonstrate a peak mode assay for the extraction of nucleic acids from E. coli cell culture. We conclude by presenting a plateau mode assay, where we use a non-focusing tracer (NFT) species to visualize the separation and perform quantitation of amino acids.

Isotachophoresis (ITP) is a robust electrokinetic separation and preconcentration technique with applications ranging from toxin detection to sample preparation. We review the physical principles of ITP and the methodology of applying this technique to two specific example applications: separation and detection of small molecules and purification of nucleic acids from cell culture lysate.

Electrokinetic techniques are a staple of microscale applications because of their unique ability to perform a variety of fluidic and electrophoretic ...

Creating Sub-50 Nm Nanofluidic Junctions in PDMS Microfluidic Chip via Self-Assembly Process of Colloidal Particles

Polydimethylsiloxane (PDMS) is the prevailing building material to make microfluidic devices due to its ease of molding and bonding as well as its transparency. Due to the softness of the PDMS material, however, it is challenging to use PDMS for building nanochannels. The channels tend to collapse easily during plasma bonding. In this paper, we present an evaporation-driven self-assembly method of silica colloidal nanoparticles to create nanofluidic junctions with sub-50 nm pores between two microchannels. The pore size as well as the surface charge of the nanofluidic junction is tunable simply by changing the colloidal silica bead size and surface functionalization outside of the assembled microfluidic device in a vial before the self-assembly process. Using the self-assembly of nanoparticles with a bead size of 300 nm, 500 nm, and 900 nm, it was possible to fabricate a porous membrane with a pore size of ~45 nm, ~75 nm and ~135 nm, respectively. Under electrical potential, this nanoporous membrane initiated ion concentration polarization (ICP) acting as a cation-selective membrane to concentrate DNA by ~1,700 times within 15 min. This non-lithographic nanofabrication process opens up a new opportunity to build a tunable nanofluidic junction for the study of nanoscale transport processes of ions and molecules inside a PDMS microfluidic chip.

We propose a simple self-assembly technique of silica colloidal nanoparticles to create a nanofluidic junction between two microchannels in polydimethylsiloxane (PDMS). Using this technique, a nanoporous bead membrane with a pore size down to ~45 nm was built inside a microchannel and applied to electrokinetic preconcentration of DNA samples.

Polydimethylsiloxane (PDMS) is the prevailing building material to make microfluidic devices due to its ease of molding and bonding as well as its ...

Engineering

A Microfluidic Chip for the Versatile Chemical Analysis of Single Cells

We present a microfluidic device that enables the quantitative determination of intracellular biomolecules in multiple single cells in parallel. For this purpose, the cells are passively trapped in the middle of a microchamber. Upon activation of the control layer, the cell is isolated from the surrounding volume in a small chamber. The surrounding volume can then be exchanged without affecting the isolated cell. However, upon short opening and closing of the chamber, the solution in the chamber can be replaced within a few hundred milliseconds. Due to the reversibility of the chambers, the cells can be exposed to different solutions sequentially in a highly controllable fashion, e.g. for incubation, washing, and finally, cell lysis. The tightly sealed microchambers enable the retention of the lysate, minimize and control the dilution after cell lysis. Since lysis and analysis occur at the same location, high sensitivity is retained because no further dilution or loss of the analytes occurs during transport. The microchamber design therefore enables the reliable and reproducible analysis of very small copy numbers of intracellular molecules (attomoles, zeptomoles) released from individual cells. Furthermore, many microchambers can be arranged in an array format, allowing the analysis of many cells at once, given that suitable optical instruments are used for monitoring. We have already used the platform for proof-of-concept studies to analyze intracellular proteins, enzymes, cofactors and second messengers in either relative or absolute quantifiable manner.

In this article we present a microfluidic chip for single cell analysis. It allows the quantification of intracellular proteins, enzymes, cofactors, and second messengers by means of fluorescent assays or immunoassays.&#160;

We present a microfluidic device that enables the quantitative determination of intracellular biomolecules in multiple single cells in parallel. For this ...

Microfluidic Chips for In Situ Crystal X-ray Diffraction and In Situ Dynamic Light Scattering for Serial Crystallography

This protocol describes fabricating microfluidic devices with low X-ray background optimized for goniometer based fixed target serial crystallography. The devices are patterned from epoxy glue using soft lithography and are suitable for in situ X-ray diffraction experiments at room temperature. The sample wells are lidded on both sides with polymeric polyimide foil windows that allow diffraction data collection with low X-ray background. This fabrication method is undemanding and inexpensive. After the sourcing of a SU-8 master wafer, all fabrication can be completed outside of a cleanroom in a typical research lab environment. The chip design and fabrication protocol utilize capillary valving to microfluidically split an aqueous reaction into defined nanoliter sized droplets. This loading mechanism avoids the sample loss from channel dead-volume and can easily be performed manually without using pumps or other equipment for fluid actuation. We describe how isolated nanoliter sized drops of protein solution can be monitored in situ by dynamic light scattering to control protein crystal nucleation and growth. After suitable crystals are grown, complete X-ray diffraction datasets can be collected using goniometer based in situ fixed target serial X-ray crystallography at room temperature. The protocol provides custom scripts to process diffraction datasets using a suite of software tools to solve and refine the protein crystal structure. This approach avoids the artefacts possibly induced during cryo-preservation or manual crystal handling in conventional crystallography experiments. We present and compare three protein structures that were solved using small crystals with dimensions of approximately 10-20 &#181;m grown in chip. By crystallizing and diffracting in situ, handling and hence mechanical disturbances of fragile crystals is minimized. The protocol details how to fabricate a custom X-ray transparent microfluidic chip suitable for in situ serial crystallography. As almost every crystal can be used for diffraction data collection, these microfluidic chips are a very efficient crystal delivery method.

Microfluidic Chips for In Situ Crystal X-ray Diffraction and In Situ Dynamic Light Scattering for Serial Crystallography

This protocol describes in detail how to fabricate and operate microfluidic devices for X-ray diffraction data collection at room temperature. Additionally, it describes how to monitor protein crystallization by dynamic light scattering and how to process and analyze obtained diffraction data.

This protocol describes fabricating microfluidic devices with low X-ray background optimized for goniometer based fixed target serial crystallography. The ...

Chemistry

A Microfluidic-based Electrochemical Biochip for Label-free DNA Hybridization Analysis

Miniaturization of analytical benchtop procedures into the micro-scale provides significant advantages in regards to reaction time, cost, and integration of pre-processing steps. Utilizing these devices towards the analysis of DNA hybridization events is important because it offers a technology for real time assessment of biomarkers at the point-of-care for various diseases. However, when the device footprint decreases the dominance of various physical phenomena increases. These phenomena influence the fabrication precision and operation reliability of the device. Therefore, there is a great need to accurately fabricate and operate these devices in a reproducible manner in order to improve the overall performance. Here, we describe the protocols and the methods used for the fabrication and the operation of a microfluidic-based electrochemical biochip for accurate analysis of DNA hybridization events. The biochip is composed of two parts: a microfluidic chip with three parallel micro-channels made of polydimethylsiloxane (PDMS), and a 3 x 3 arrayed electrochemical micro-chip. The DNA hybridization events are detected using electrochemical impedance spectroscopy (EIS) analysis. The EIS analysis enables monitoring variations of the properties of the electrochemical system that are dominant at these length scales. With the ability to monitor changes of both charge transfer and diffusional resistance with the biosensor, we demonstrate the selectivity to complementary ssDNA targets, a calculated detection limit of 3.8 nM, and a 13% cross-reactivity with other non-complementary ssDNA following 20 min&#160;of incubation. This methodology can improve the performance of miniaturized devices by elucidating on the behavior of diffusion at the micro-scale regime and by enabling the study of DNA hybridization events.

We present a microfluidic-based electrochemical biochip for DNA hybridization detection. Following ssDNA probe functionalization, the specificity, sensitivity, and detection limit are studied with complementary and non-complementary ssDNA targets. Results illustrate the influence of the DNA hybridization events on the electrochemical system, with a detection limit of 3.8 nM.

Miniaturization of analytical benchtop procedures into the micro-scale provides significant advantages in regards to reaction time, cost, and integration ...

A Microfluidic Platform for Precision Small-volume Sample Processing and Its Use to Size Separate Biological Particles with an Acoustic Microdevice

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. However, to achieve robust sample manipulation it is necessary to address device integration with the macroscale environment. To realize repeatable, sensitive particle separation with microfluidic devices, this protocol presents a complete automated and integrated microfluidic platform that enables precise processing of 0.15&#8211;1.5 ml samples using microfluidic devices. Important aspects of this system include modular device layout and robust fixtures resulting in reliable and flexible world to chip connections, and fully-automated fluid handling which accomplishes closed-loop sample collection, system cleaning and priming steps to ensure repeatable operation. Different microfluidic devices can be used interchangeably with this architecture. Here we incorporate an acoustofluidic device, detail its characterization, performance optimization, and demonstrate its use for size-separation of biological samples. By using real-time feedback during separation experiments, sample collection is optimized to conserve and concentrate sample. Although requiring the integration of multiple pieces of equipment, advantages of this architecture include the ability to process unknown samples with no additional system optimization, ease of device replacement, and precise, robust sample processing.

This protocol describes a system architecture for performing automated small volume (0.15&#8211;1.5 ml) particle separations using a microfluidic device, and discusses methods to optimize acoustofluidic device performance and operation.

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. ...

Fabrication of a Dipole-assisted Solid Phase Extraction Microchip for Trace Metal Analysis in Water Samples

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water samples. A brief overview of the evolution of chip-based SPE techniques is provided. This is followed by an introduction to specific polymeric materials and their role in SPE. To develop an innovative dipole-assisted SPE technique, a chlorine (Cl)-containing SPE functionality was implanted into a poly(methyl methacrylate) (PMMA) microchip. Herein, diverse analytical techniques including contact angle analysis, Raman spectroscopic analysis, and laser ablation-inductively coupled plasma-mass spectrometry (LA-ICP-MS) analysis were employed to validate the utility of the implantation protocol of the C-Cl moieties on the PMMA. The analytical results of the X-ray absorption near-edge structure (XANES) analysis also demonstrated the feasibility of the Cl-containing PMMA used as an extraction medium by virtue of the dipole-ion interactions between the highly electronegative C-Cl moieties and the positively charged metal ions.

The fabrication protocol of a dipole-assisted solid phase extraction microchip for the trace metal analysis is presented.

This paper describes a fabrication protocol for a dipole-assisted solid phase extraction (SPE) microchip available for trace metal analysis in water ...

Fast Enzymatic Processing of Proteins for MS Detection with a Flow-through Microreactor

The vast majority of mass spectrometry (MS)-based protein analysis methods involve an enzymatic digestion step prior to detection, typically with trypsin. This step is necessary for the generation of small molecular weight peptides, generally with MW &#60; 3,000-4,000 Da, that fall within the effective scan range of mass spectrometry instrumentation. Conventional protocols involve O/N enzymatic digestion at 37 &#186;C. Recent advances have led to the development of a variety of strategies, typically involving the use of a microreactor with immobilized enzymes or of a range of complementary physical processes that reduce the time necessary for proteolytic digestion to a few minutes (e.g., microwave or high-pressure). In this work, we describe a simple and cost-effective approach that can be implemented in any laboratory for achieving fast enzymatic digestion of a protein. The protein (or protein mixture) is adsorbed on C18-bonded reversed-phase high performance liquid chromatography (HPLC) silica particles preloaded in a capillary column, and trypsin in aqueous buffer is infused over the particles for a short period of time. To enable on-line MS detection, the tryptic peptides are eluted with a solvent system with increased organic content directly in the MS ion source. This approach avoids the use of high-priced immobilized enzyme particles and does not necessitate any aid for completing the process. Protein digestion and complete sample analysis can be accomplished in less than ~3 min and ~30 min, respectively.

A quick protocol for proteolytic digestion with an in-house built flow-through tryptic microreactor coupled to an electrospray ionization (ESI) mass spectrometer is presented. The fabrication of the microreactor, the experimental setup and the data acquisition process are described.

The vast majority of mass spectrometry (MS)-based protein analysis methods involve an enzymatic digestion step prior to detection, typically with trypsin. ...

A Microfluidic Chip for ICPMS Sample Introduction

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