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Lens stacks can require adjustments made during the assembly into the endoscope tip and along the entire path from the objective lens to the image sensor, but must be sealed to keep fluids out of the optical path.

To design and construct a practical chip-on-tip device, the camera and the LEDs need to be mechanically mounted within a tip. In the case of a medical endoscope, it is typically necessary to keep the girth of the tip as small as possible.

This means the tip will have very small features and thin walls. Endoscopy equipment. Image courtesy of the National Cancer Institute.

However, injection molding parts this small requires specialized micro molding techniques and practices. Successful micro molding begins with the component part design, through to tool design, and finally to proper molding equipment and processing parameters.

The attributes of conventional molding processes and equipment have limitations that can preclude effective molding of micro-scale parts. Large barrel sizes create problems with material residence times.

Managing small shot sizes with the precision needed is not possible for very small shot sizes using large equipment and will result in short shots or flash from overpacking.

Part design can dictate mold construction. Without care and robust design review practices, it is possible to create part details that would function perfectly in the finished part but will result in poor mold robustness or complexity of construction.

This can result in making the part more difficult or even impossible to produce. Image sensors have various analog and digital formats for data output, and some are more sensitive than others.

The data moving from the image sensor to its destination is susceptible to corruption or distortion from both internal e.

Preserving signal integrity begins with the design of the circuit to drive the LEDs. By employing PWM, the pulse magnitude and duration may be varied to employ a variable intensity i.

PWM can also drive the LEDs with a peak current higher than the maximum continuous current to achieve a higher lumen output.

Analog signals vs. PWM signals. Achieving proper shielding in extremely small chip-on-tip designs can be accomplished with micro-coax wiring and flex circuits that employ shielding layers to emulate a coax shield.

The challenge of preserving signal integrity is exacerbated when the distance from the sensor to the display or storage device is increased, reducing signal strength and multiplying opportunities for interference.

Solutions in these instances include amplifier circuits and low-loss conductors. Use of low-loss conductors becomes a compromise of increasing conductor size to the limits of the available space.

Solutions can be created with alternate geometries to take advantage of the space available. Two examples include employing flat wire and printing conductors directly onto enclosures and cannulae.

Of course, the interconnection needs to be environmentally sealed to prevent shorting and other influences of exposure to fluids.

In medical applications, this typically means saline solution, bodily fluids, and other substances. Successful sealing begins at the design stage by providing appropriate features for the sealing system to be employed.

This may mean including wells designed to accept and contain potting compounds, for example. Using properly selected potting compounds is one of the most practical ways to seal items of this scale.

Mechanical sealing systems using gaskets, for example, would increase the quantity of micro-scale components and further complicate proper assembly.

When selecting appropriate potting compounds, there are several factors to consider. Potting materials come in several different chemistries. Single-part chemistries are typically designed to cure quickly using heat or UV light or over time at room temperature.

Room temperature curing materials experience viscosity changes over time, which adds to process variability and pot life i. Additional complications can arise from cure times that can extend over dozens of hours.

In these cases, sub-assemblies must be carefully fixtured to prevent movement or displacement during the curing process. However, this approach results in high yield-risk, inefficient use of space, difficult handling, and a great deal of work in process WIP delays.

UV curing materials cure very quickly, typically in a matter of seconds, but require that the entirety of the dispensed material can be exposed to the UV light source and not be in a shadowed area.

As such, UV materials are typically translucent. However, translucent materials can present a significant limitation when seeking to block areas of the image sensor from extraneous light.

Environmental protections that extend beyond potting processes are also required. For instance, the image sensor, LEDs, and electronics in chip-on-tip endoscopes need to reliably withstand medical sterilization and sanitation processes.

This could include the high heat and moisture of an autoclave, ethylene oxide, glutaraldehyde, vaporized hydrogen peroxide, irradiation, and iodine.

Falanga, and P. There have been many reports on microfluidic devices for cell culture having upper and lower microchannels separated by a thin PDMS membrane.

In these devices, the lower channel often interferes with the microscopic observation of cells cultured in the upper channel. To avoid interference, a microdevice with a detachable lower channel was developed.

Mix the elastomer and curing agent at a mass ratio. De-gas the mixture under vacuum until no bubbles remain 20 min. Punch the inlet and outlet holes at both the ends of the upper channel with a 2-mm biopsy punch.

Remove the PMMA sheet, and punch a hole to connect the sheet with the lower channel by using a 1-mm biopsy punch from the membrane side.

Place the lower sheet on the coated glass slide Fig. Peel off the lower sheet from the glass slide and place the glue-coated surface of the sheet on the PDMS membrane Fig.

After 30 min of incubation, bond the lower sheet to a cover slip by plasma bonding. Introduce a cell suspension into the upper microchannel, which is manually precoated with 0.

Remove the lower sheet from the device carefully Fig. Place the rest of the device on a cover slip for observation with an inverted microscope Fig.

The cell culture channel upper is filled with water containing a red food color, while the lower channel is filled with water containing a blue food color.

Phase contrast images of cells e before and f after detachment of the lower sheet. We developed a microfluidic device with a detachable lower microchannel.

It is important that different bonding techniques be used for each side of the PDMS membrane. If the lower channel is filled with air and the device is incubated in a CO 2 incubator, dew condensation is often observed in the lower channel when the device is taken out from the incubator.

The condensation in the lower channel makes observation difficult Fig. This problem was solved with the detachable device.

The demand for microfluidics has steadily increased, due in part to the growing popularity of point-of-care devices [1]. Often, microfluidic chips are fabricated in thermoplastics [1].

Thermoplastics are synthetic polymers that have gained popularity due to their ability to be molded into complex structures [3, 4].

They are often used as a safer and cheaper alternative to glass [3, 4]. However, proper sealing of these devices proves challenging, especially in the field of medical testing, where the demand for reliable devices is high.

For example, pressure-sensitive adhesives, common sealants, can limit the size of microfluidic channels; some adhesive can exhibit reactive groups that interfere with analytical processes that run on the chip [5].

Hence, a method of sealing that is free from the aforementioned limitations is needed. Here, a solvent-based method is presented.

Polymethylmethacrylate PMMA , a thermoplastic, exhibits softening at temperatures above its glass transition temperature T g returning to its original state when cooled.

This transition introduces several direct bonding options [6]. The pressure required for bonding even at this temperature is fairly high.

This can lead to imperfections in the channel dimensions, as the bulk of the material softens. The application of a weak solvent decreases T g only for the surface of the plastic, thus reducing the required temperature and pressure for the process.

The decreased pressure reduces the possibility of channel deformation. Furthermore, as the solvent-induced softening is limited only to the surface the first few microns , the deeper channel structures are not affected.

Hence, a direct solvent bonding method allows for an adhesive-free bonding and avoids a temperature-induced deformation.

As a bonus, the mechanical properties of the bond are greatly enhanced [7]. It is worth noting that this approach is valid for microfluidic devices with channel depths greater than microns, typically for devices produced by a direct laser etching.

Another advantage of this technique is that it results in the production of sterile devices when the weak solvent is ethanol.

They can be manufactured quickly using basic equipment found in any laboratory [7]. Bonding setup. A Alignment manifold B 3 wooden pins are used to keep the layers from moving.

Email: saifullah. The subject of droplet microfluidics has grown in importance among researchers in chemistry, physics and biology, hence it has found applications in drug delivery, encapsulation, single-cell analysis, pickering-emulsion and phase-separation.

For generating monodisperse droplets, various methods have been employed in constructing microfluidic devices. Small channel-diameters attained by clean-room soft lithography is the most precise technique for fabricating microfluidic devices.

Therefore, the cost and special clean-room training restricts its wide-spread application. Recently, a rapid prototyping technique for microfluidics has been reported by employing laser-patterned tape 4 This technique relies on computer-controlled CO 2 laser beam.

This work was further simplified by manual razor patterned tape-based prototyping for patterning mammalian cells.

Hence, our approach may well serve as one of the simplest approaches to fabricate droplet microfluidic generators.

Figure 1 outlines the prototyping procedure. Prototyping begins by attaching adhesive tape on a flat glass substrate.

With a sharp razor-blade, the tape is cut into fine parallel strips. Next the tape is removed from the regions outside the fine strips.

The junction is pressed gently to ensure the strips are well attached. These adhering strips of tape serve as a master for PDMS-based replica casting.

A mixture of PDMS silicone elastomer base and a curing agent in ratio is poured on top of the master within a plastic petri dish.

Cured PDMS replica is then cut and peeled-off from the master. The master can be used repeatedly to fabricate multiple copies of the PDMS replica by following the afore-mentioned steps.

Inlet and outlet holes are drilled through PDMS replica, which is then bonded on a glass substrate, after both replica and glass has been exposed to oxygen plasma.

The technique is easily extended to fabricate T-junction or double T-junction prototypes Figure 1h and i. As the outer flow-rate is increased, the regime is found to shift from dripping at lower flow-rate to jetting at higher flow-rate Figure 2 c.

For lowest flow-rate, the aqueous-phase breaks into elongated plugs, while at higher flow-rates regular drops are pinched off.

Figure 2b shows the droplet-size as a function of Ca. Rapid Prototyping of Microfluidic Systems in Poly dimethyl siloxane.

Rapid prototyping of microfluidic systems using a laser-patterned tape J. Adhesive-tape soft lithography for patterning mammalian cells: application to wound-healing assays.

BioTechniques, , 53 — Greiner, A. Microfluidic devices are used for many different types of experiments across the medical, ecological and evolutionary disciplines Park et al.

For example, microfluidic devices for microbial experiments require inoculation into smaller chambers that simulate natural microbial environments such as porous soils Or et al.

These devices often involve complicated pump setups and irreversible seals. We developed a technique that requires only common lab equipment and makes the device reusable while also allowing the microbes to grow undisturbed based on Tekwa et al.

Here, we provide a detailed guide for the assembly and the previously undocumented non-destructive disassembly of polydimethylsiloxane PDMS experimental devices to recover microbes in situ , which can then be plated for relative counts and further molecular analyses of population changes.

This is complemented by videos for each step. Figure 1: Microfluidic device containing 14 habitats on an elastomer PDMS layer pressed onto a 60mm x 24mm glass cover slip.

This device is used to test the effects of habitat patchiness on microbe dynamics. Habitats were dyed blue for visualization.

For more information see Tekwa et al. Figure 4. View of an inoculated and incubated device, looking through the bottom of a petri dish.

The recovery technique can be used to estimate relative proportions of different types of microbes e. Unlike in Tekwa et al. These videos go through the specific procedure that we used to perform experiments on competition and cooperation in Pseudomonas aeruginosa and may be useful in determining specific amounts of media, growth times, etc.

Cho, H. Self-organization in high-density bacterial colonies: efficient crowd control. PLoS biology , 5 11 , e Connell, J.

Proceedings of the National Academy of Sciences , 46 , Folkesson, A. Adaptation of Pseudomonas aeruginosa to the cystic fibrosis airway: an evolutionary perspective.

Nature reviews. Microbiology , 10 12 , Hol, F. Zooming in to see the bigger picture: Microfluidic and nanofabrication tools to study bacteria.

Science , , Keymer, J. Computation of mutual fitness by competing bacteria. Proceedings of the National Academy of Sciences , 51 , Or, D.

Physical constraints affecting bacterial habitats and activity in unsaturated porous media — a review. Advances in Water Resources , 30 6 , Park, S.

Motion to form a quorum. Tekwa, E. Patchiness in a microhabitat chip affects evolutionary dynamics of bacterial cooperation. Lab on a Chip , 15 18 , Defector clustering is linked to cooperation in a pathogenic bacterium.

In review. Paris 06, Paris, France. Glass is a versatile surface for chemical treatments, and it still is by far the most used substrate for surface engineering e.

For cell culture on such substrates, glass-bottom culture dishes are desired to keep over the cells well defined medium volumes, and to protect the cells from contamination and medium evaporation.

Moreover, they are optically better suited for microscopy observation than polystyrene dishes routinely used for cell culture.

Although glass-bottom culture dishes are commercially available e. In this Tip, we describe an easier way than a previous Tip 1 to transform a polystyrene culture dish into a glass-bottom one, while preserving the possibility to apply to the glass any treatment before its assembly into a dish.

Note that in this method, the body of the culture dish will be upside down and the lid is thus no longer lifted above the dish opening by lid stoppers.

However, the gas exchange through the gap between the body and its lid seems to be enough to culture cells healthily in this dish.

In summary, this Tip provides a low-cost and rapid solution for cell culture in a microfluidic device or on an engineered surface directly in a culture dish, suited for a long-term live cell imaging.

Why is it useful? Modern microfluidic devices can incorporate channels of different heights to fulfill their designed function.

Examples include hydrodynamic focusing [1], cell traps [2], and chambers that isolate cellular components [3]. These devices are fabricated from a multilayer SU-8 photoresist master mold.

Each layer height requires a separate set of photolithographic steps, including photoresist spin, photomask alignment, exposure, and bakes, followed by a development step at the end to reveal the 3D resist pattern.

They are indispensable tools for creating multilayer patterns with accurate registration, but while available in cleanrooms at many research universities, their substantial expense may place them out of reach of teaching institutions and individual laboratories.

In contrast, single-layer microfluidics can be prepared using an inexpensive UV light source, or even a self-made one [4].

In principle, manual photomask alignment could be made under a microscope, then brought to the UV source, yet this poses several complications. First, alignment features can be very difficult to see using inexpensive microscopes or stereoscopes, especially in thin SU8 layers, due to poor contrast between exposed and unexposed regions before development.

Second, misalignment can occur during movement to the exposure system. In this tip, we present a method for manual alignment of multiple transparency photomasks.

These accuracies are within required tolerances of many multilayer designs Figure 4b. In many cases, minor design alternatives can relax alignment tolerances, such as in a trap design containing a thin horizontal channel that allows fluid bypass but captures larger objects Figure 4c.

Furthermore, after alignment marks are cut, no microscope is needed at all during the photolithography process, speeding the fabrication of multiple masters.

Lab-on-a-chip LOC devices significantly contribute different disciplines of science. Polydimethylsiloxane PDMS is one of the main materials, which is widely used for the fabrication of biological LOCs, due to its biocompatibility and ease of use.

However, PDMS and some other polymeric materials are intrinsically water repellant or hydrophobic , which results in difficulties in loading and operating LOCs.

The eminent consequence of hydrophobicity in LOCs for biological systems is the entrapment of air bubbles in microfluidic channels.

Although the oxygen plasma treatment of PDMS reduces the surface hydrophobicity for a certain period of time, the hydrophilic property of PDMS vanishes over time 1.

The persistent problem of bubbles in the microfluidics led to several studies conducted to overcome it. Some of these solutions suggested implementing bubble traps 2 , 3 , surface treatment of LOCs through hydrophilic coatings 4 , and using actively controlled bubble removal systems 5 , 6.

Although the aforementioned design complexities are introduced to LOCs in order to reduce the clogging problem caused by the bubbles, these modifications also result in higher production cost, complex operation, and long device preparation time.

In many single-cell experiments without losing or damaging the rare cells, these cells needs to be introduce into the LOCs.

Here, we present a simple method that enables loading a small number of cells without introducing bubbles in the microfluidics channels.

Thus, the inner surface of microfluidic channels will be disinfected and the fluid flow will be tested within the micro channels as it is applied in many other protocols for LOCs 7 , 8.

Step 3: Gently apply pressure pressing the pipetman to force ethanol solution flow through the micro channels and cavities of the PDMS device.

Take care to avoid applying negative pressure from the outlet-pipet tip, which might create air leakage through the pipet connections. The positive pressure will facilitate removal of the air bubbles via dissolving them.

Step 4: After flushing the chip with ethanol solution, inspect the chip to ensure bubble removal. In case of air bubbles, repeat the steps 2 and 3. Step 5: Fill the syringe 10 ml with medium or phosphate buffered saline PBS.

Take care to remove the air bubbles inside the syringe, mount and lock the needle on the syringe. Then, flow medium through the needle too make sure the needle is full of medium without any bubble.

Insert the needle in the inlet-pipet tip; gently apply positive pressure to replace the ethanol with medium. Next, collect the excess medium from the outlet-pipet tip.

Step 6: Fill the inlet pipet with fresh medium in such a way that due to certain height h between the levels of the medium in the inlet and outlet pipet tips, very slow medium flow will be established inside the micro-channels.

Step 7: Load your cells into the Hamilton syringe and take care to ensure that there is no air bubble inside its needle and syringe.

Insert the needle of the Hamilton syringe into the inlet-pipet tip as explained in Step 5, Figure 1. Introduce the cells via applying gentle positive pressure to the syringe.

Established flow streams in the PDMS chip will deliver the released cells to the desired positions in the chip.

Flow rate can be arranged adjusting the applied positive pressure and amount of medium collected in the inlet and outlet pipet tips. The excess supernatant from the outlet-pipet tip can be collected, and fresh medium can be supplied through the inlet-pipet tip during the experiment.

Chips and Tips RSS. Chips and Tips. By Liz Bowley. Do you have problems with bubble formation when injecting your sample? Or do you have your own tricks to overcome problems like these?

By Keir Hollingsworth. Jonathan Tjong 1 , Alyne G. Teixeira 1 and John P. What Do I Need? After cleaning and post-curing of the 3D-printed mold in the UV lightbox, place the mold in the container and add enough solvent to submerge the part.

Seal the container and leave on a shaker table for 24 hours. Discard the old solvent and add new solvent. Seal and agitate for another 24 hours.

Remove the part from the solvent and allow to air dry at room temperature. What else should I know? References B9Creations. Cutting the cords: Two paths to well-plate microfluidics 25 Mar By Sian Carrington, Editorial Assistant.

A second life for old electronic parts: a spin coater for microfluidic applications 25 Apr What do I need? Assembling of spin coater 1.

Remove the fan from an old pc or mac, if are particularly posh Fig. Development of a cell culture microdevice with a detachable channel for clear observation 21 Feb PDMS molding Mix the elastomer and curing agent at a mass ratio.

By Harriet Riley, Development Editor. Cheong b and S. Panels h and i show razor patterned tape-based T-junction and double T-junction prototypes, respectively.

Reference [1] Qin, D. Soft Lithography. Why is this useful? Fill and empty the container with water 10 times in order to rinse the devices.

Lastly, fill the container with filtered water, seal with tin foil and autoclave in order to sterilize the devices. Place the devices with features facing up on the lid of a petri dish and place a small amount i.

Using sterile tweezers, pick up the device and place face down into the centre of a petri dish or cover glass, sealing it to the surface by pushing on the back with gloved fingers repeatedly, using a kimwipe to wick excess liquid away from the side.

Then surround, but not touch, the device with kimwipes soaked in filtered water Fig. Place upright in incubator for the desired amount of time.

The experiment can now proceed untouched for up to 24 hours see Supplementary video. Device with bacteria droplets, 1 droplet per habitat.

Recovering from the devices perform in a BSC : Open petri dish, carefully remove and discard kimwipes and then use sterile tweezers to gently unseal the device and place face up in the lid of the petri dish.

By around 12 hours, the spaces between the habitats will be void of liquid from PDMS absorption, preventing microbes from being mixed across chambers during disassembly.

Habitats that have dried out will appear white Fig. Dip sterile inoculating loop into an eppendorf with PBS, then use that loop to scrape one of the habitats Fig.

Dip that inoculating loop back into the eppendorf media again, which can then be grown overnight for further analyses such as plating for relative cell count if there are different strains and other molecular analyses.

Repeat for the rest of the habitats that you are interested in, using new inoculating loops. Figure 5. Recovering bacteria from a habitat in the disassembled PDMS device.

Links to Videos These videos go through the specific procedure that we used to perform experiments on competition and cooperation in Pseudomonas aeruginosa and may be useful in determining specific amounts of media, growth times, etc.

References Cho, H. Rapid and easy fabrication of glass-bottom culture dishes for long-term live cell imaging 25 Jul Place a polystyrene culture dish upside down on a surface and hit a few times the center of the dish bottom with the grip of a large screw driver Fig.

The bottom should fall easily with the success rate around 9 over Avoid breaking the dish wall by hitting the bottom too strongly.

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Connell, J. Proceedings of the National Academy of Sciences , 46 , Folkesson, A. Adaptation of Pseudomonas aeruginosa to the cystic fibrosis airway: an evolutionary perspective.

Nature reviews. Microbiology , 10 12 , Hol, F. Zooming in to see the bigger picture: Microfluidic and nanofabrication tools to study bacteria.

Science , , Keymer, J. Computation of mutual fitness by competing bacteria. Proceedings of the National Academy of Sciences , 51 , Or, D.

Physical constraints affecting bacterial habitats and activity in unsaturated porous media — a review. Advances in Water Resources , 30 6 , Park, S.

Motion to form a quorum. Tekwa, E. Patchiness in a microhabitat chip affects evolutionary dynamics of bacterial cooperation. Lab on a Chip , 15 18 , Defector clustering is linked to cooperation in a pathogenic bacterium.

In review. Paris 06, Paris, France. Glass is a versatile surface for chemical treatments, and it still is by far the most used substrate for surface engineering e.

For cell culture on such substrates, glass-bottom culture dishes are desired to keep over the cells well defined medium volumes, and to protect the cells from contamination and medium evaporation.

Moreover, they are optically better suited for microscopy observation than polystyrene dishes routinely used for cell culture.

Although glass-bottom culture dishes are commercially available e. In this Tip, we describe an easier way than a previous Tip 1 to transform a polystyrene culture dish into a glass-bottom one, while preserving the possibility to apply to the glass any treatment before its assembly into a dish.

Note that in this method, the body of the culture dish will be upside down and the lid is thus no longer lifted above the dish opening by lid stoppers.

However, the gas exchange through the gap between the body and its lid seems to be enough to culture cells healthily in this dish. In summary, this Tip provides a low-cost and rapid solution for cell culture in a microfluidic device or on an engineered surface directly in a culture dish, suited for a long-term live cell imaging.

Why is it useful? Modern microfluidic devices can incorporate channels of different heights to fulfill their designed function.

Examples include hydrodynamic focusing [1], cell traps [2], and chambers that isolate cellular components [3]. These devices are fabricated from a multilayer SU-8 photoresist master mold.

Each layer height requires a separate set of photolithographic steps, including photoresist spin, photomask alignment, exposure, and bakes, followed by a development step at the end to reveal the 3D resist pattern.

They are indispensable tools for creating multilayer patterns with accurate registration, but while available in cleanrooms at many research universities, their substantial expense may place them out of reach of teaching institutions and individual laboratories.

In contrast, single-layer microfluidics can be prepared using an inexpensive UV light source, or even a self-made one [4]. In principle, manual photomask alignment could be made under a microscope, then brought to the UV source, yet this poses several complications.

First, alignment features can be very difficult to see using inexpensive microscopes or stereoscopes, especially in thin SU8 layers, due to poor contrast between exposed and unexposed regions before development.

Second, misalignment can occur during movement to the exposure system. In this tip, we present a method for manual alignment of multiple transparency photomasks.

These accuracies are within required tolerances of many multilayer designs Figure 4b. In many cases, minor design alternatives can relax alignment tolerances, such as in a trap design containing a thin horizontal channel that allows fluid bypass but captures larger objects Figure 4c.

Furthermore, after alignment marks are cut, no microscope is needed at all during the photolithography process, speeding the fabrication of multiple masters.

Lab-on-a-chip LOC devices significantly contribute different disciplines of science. Polydimethylsiloxane PDMS is one of the main materials, which is widely used for the fabrication of biological LOCs, due to its biocompatibility and ease of use.

However, PDMS and some other polymeric materials are intrinsically water repellant or hydrophobic , which results in difficulties in loading and operating LOCs.

The eminent consequence of hydrophobicity in LOCs for biological systems is the entrapment of air bubbles in microfluidic channels.

Although the oxygen plasma treatment of PDMS reduces the surface hydrophobicity for a certain period of time, the hydrophilic property of PDMS vanishes over time 1.

The persistent problem of bubbles in the microfluidics led to several studies conducted to overcome it.

Some of these solutions suggested implementing bubble traps 2 , 3 , surface treatment of LOCs through hydrophilic coatings 4 , and using actively controlled bubble removal systems 5 , 6.

Although the aforementioned design complexities are introduced to LOCs in order to reduce the clogging problem caused by the bubbles, these modifications also result in higher production cost, complex operation, and long device preparation time.

In many single-cell experiments without losing or damaging the rare cells, these cells needs to be introduce into the LOCs. Here, we present a simple method that enables loading a small number of cells without introducing bubbles in the microfluidics channels.

Thus, the inner surface of microfluidic channels will be disinfected and the fluid flow will be tested within the micro channels as it is applied in many other protocols for LOCs 7 , 8.

Step 3: Gently apply pressure pressing the pipetman to force ethanol solution flow through the micro channels and cavities of the PDMS device.

Take care to avoid applying negative pressure from the outlet-pipet tip, which might create air leakage through the pipet connections.

The positive pressure will facilitate removal of the air bubbles via dissolving them. Step 4: After flushing the chip with ethanol solution, inspect the chip to ensure bubble removal.

In case of air bubbles, repeat the steps 2 and 3. Step 5: Fill the syringe 10 ml with medium or phosphate buffered saline PBS.

Take care to remove the air bubbles inside the syringe, mount and lock the needle on the syringe. Then, flow medium through the needle too make sure the needle is full of medium without any bubble.

Insert the needle in the inlet-pipet tip; gently apply positive pressure to replace the ethanol with medium. Next, collect the excess medium from the outlet-pipet tip.

Step 6: Fill the inlet pipet with fresh medium in such a way that due to certain height h between the levels of the medium in the inlet and outlet pipet tips, very slow medium flow will be established inside the micro-channels.

Step 7: Load your cells into the Hamilton syringe and take care to ensure that there is no air bubble inside its needle and syringe.

Insert the needle of the Hamilton syringe into the inlet-pipet tip as explained in Step 5, Figure 1. Introduce the cells via applying gentle positive pressure to the syringe.

Established flow streams in the PDMS chip will deliver the released cells to the desired positions in the chip. Flow rate can be arranged adjusting the applied positive pressure and amount of medium collected in the inlet and outlet pipet tips.

The excess supernatant from the outlet-pipet tip can be collected, and fresh medium can be supplied through the inlet-pipet tip during the experiment.

Chips and Tips RSS. Chips and Tips. By Liz Bowley. Do you have problems with bubble formation when injecting your sample? Or do you have your own tricks to overcome problems like these?

By Keir Hollingsworth. Jonathan Tjong 1 , Alyne G. Teixeira 1 and John P. What Do I Need? After cleaning and post-curing of the 3D-printed mold in the UV lightbox, place the mold in the container and add enough solvent to submerge the part.

Seal the container and leave on a shaker table for 24 hours. Discard the old solvent and add new solvent. Seal and agitate for another 24 hours.

Remove the part from the solvent and allow to air dry at room temperature. What else should I know? References B9Creations. Cutting the cords: Two paths to well-plate microfluidics 25 Mar By Sian Carrington, Editorial Assistant.

A second life for old electronic parts: a spin coater for microfluidic applications 25 Apr What do I need? Assembling of spin coater 1.

Remove the fan from an old pc or mac, if are particularly posh Fig. Development of a cell culture microdevice with a detachable channel for clear observation 21 Feb PDMS molding Mix the elastomer and curing agent at a mass ratio.

By Harriet Riley, Development Editor. Cheong b and S. Panels h and i show razor patterned tape-based T-junction and double T-junction prototypes, respectively.

Reference [1] Qin, D. Soft Lithography. Why is this useful? Fill and empty the container with water 10 times in order to rinse the devices.

Lastly, fill the container with filtered water, seal with tin foil and autoclave in order to sterilize the devices. Place the devices with features facing up on the lid of a petri dish and place a small amount i.

Using sterile tweezers, pick up the device and place face down into the centre of a petri dish or cover glass, sealing it to the surface by pushing on the back with gloved fingers repeatedly, using a kimwipe to wick excess liquid away from the side.

Then surround, but not touch, the device with kimwipes soaked in filtered water Fig. Place upright in incubator for the desired amount of time.

The experiment can now proceed untouched for up to 24 hours see Supplementary video. Device with bacteria droplets, 1 droplet per habitat.

Recovering from the devices perform in a BSC : Open petri dish, carefully remove and discard kimwipes and then use sterile tweezers to gently unseal the device and place face up in the lid of the petri dish.

By around 12 hours, the spaces between the habitats will be void of liquid from PDMS absorption, preventing microbes from being mixed across chambers during disassembly.

Habitats that have dried out will appear white Fig. Dip sterile inoculating loop into an eppendorf with PBS, then use that loop to scrape one of the habitats Fig.

Dip that inoculating loop back into the eppendorf media again, which can then be grown overnight for further analyses such as plating for relative cell count if there are different strains and other molecular analyses.

Repeat for the rest of the habitats that you are interested in, using new inoculating loops. Figure 5. Recovering bacteria from a habitat in the disassembled PDMS device.

Links to Videos These videos go through the specific procedure that we used to perform experiments on competition and cooperation in Pseudomonas aeruginosa and may be useful in determining specific amounts of media, growth times, etc.

References Cho, H. Rapid and easy fabrication of glass-bottom culture dishes for long-term live cell imaging 25 Jul Place a polystyrene culture dish upside down on a surface and hit a few times the center of the dish bottom with the grip of a large screw driver Fig.

The bottom should fall easily with the success rate around 9 over Avoid breaking the dish wall by hitting the bottom too strongly. Spread uncured PDMS mixture on a flat substrate e.

Place the dish broken part up on a cover glass slide Fig. Surface treatment e. To keep humidity for on-chip cell culture, the dish can be filled with e.

Dishes with chips or micropatterns loaded with cells can be placed in a CO 2 incubator with or without further protection Fig.

Long-term live cell imaging can be performed using a stage top incubator Fig. Depending on the support type of microscopes, it might be necessary to well align the contours of the dish and the glass slide.

This can be done using cylindrical magnets 3 per dish and a ferromagnetic metal plate Fig. Reference [1] Caballero D, Samitier J, Different strategies for the fabrication of cell culture chambers for live-cell imaging studies.

Multilayer photolithography with manual photomask alignment 05 Jun Frank Benesch-Lee, Jose M. Lazaro Guevara, and Dirk R.

Equipment and supplies for photolithography: Spin coater, and UV exposure system Substrate wafer and SU-8 photoresist Small microscope e. Cut the photomasks from the transparency sheet, leaving 4 corner tabs.

Align the two masks relative to each other under the microscope Figure 1a and clip them together with a binder clip.

Ensure correct mask orientation and check alignment accuracy at multiple alignment marks across the mask. Align in the vertical direction first then rotate the masks 90 degrees to ensure accurate alignment in both horizontal and vertical directions.

Add binder clips to each corner Figure 1b , and verify alignment. Next, remove one binder clip at a time and use a straight razor blade to cut a sharp V-notch into each tab, through both masks.

Press the blade straight down to avoid shifting the alignment. Replace the binder clip, and proceed to the next corner until all 4 notches are cut Figure 1c.

Spin the first layer of SU-8 onto the wafer to the desired thickness and prebake. Attach 4 pieces of scotch tape onto the bottom of the wafer so that the sticky side faces up Figure 2a.

Position the first mask on the wafer, pressing gently to adhere it to the tape tabs. Use a fine-tip marker to trace the alignment notches Figure 2b onto the scotch tape Figure 2c.

Transfer to the UV exposure system and expose. Carefully remove the mask without detaching the scotch tape from the wafer and postbake.

Apply an additional piece of tape to cover the sticky tape tabs to protect the marker from smearing and allow smooth alignment of the next mask.

Spin coat the next photoresist layer and prebake Figure 3a. Mount the wafer onto a glass plate with a loop of scotch tape to keep it in place.

Affix the mask to the glass plate with thin mm wide pieces of tape, and adjust alignment as necessary.

Carefully transfer the glass plate with wafer and aligned photomask for exposure Figure 3c. Repeat step 3 for any additional layers.

Remove the tape tabs and develop the photoresist. Evaluate alignment accuracy under a microscope Figure 4. Conclusions: In this tip, we present a method for manual alignment of multiple transparency photomasks.

Zhi-Xiong, and Y. Journal of Micromechanics and Microengineering, Erickson, J. Journal of Neuroscience Methods, Taylor, A.

Nature Methods, Erickstad, M. Gutierrez, and A. Groisman, A low-cost low-maintenance ultraviolet lithography light source based on light-emitting diodes.

Lab on a Chip, A simple, bubble-free cell loading technique for culturing mammalian cells on lab-on-a-chip devices 28 Feb Purpose Lab-on-a-chip LOC devices significantly contribute different disciplines of science.

References Tan, S. Nguyen, Y. Chua, and T. Biomicrofluidics, Zheng, W. Wang, W. Zhang, and X. Lab on a Chip , Wang, Y.

Lee, L. Zhang, H. Jeon, J. Mendoza-Elias, T. Harvat, S. Hassan, A. Zhou, D. Eddington, and J. Biomedical Microdevices , Sims, and N.

Karlsson, J. Gazin, S. Laakso, T. Haraldsson, S. Malhotra-Kumar, M. Maki, H. Goossens, and W. Cortes, D. Tang, D. Capelluto, and I.

Sensors and Actuators B: Chemical , Benavente-Babace, A. Gallego-Perez, D. Hansford, S. Arana, E. Perez-Lorenzo, and M.

Yesilkoy, F. Ueno, B. Desbiolles, M. Grisi, Y. Sakai, B. Safety status of Chip-tip. Get more Chip-tip. Latest check 24 days ago. Countable Data Brief.

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