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Afterwards, to turn on the fan, connect the female and male pins. Using the tesa power strips, secure the substrate i.

For devices larger than the fan, use an adeguate plastic stopper to elevate the device right picture. Drip, by a micro pipette, the liquid containing the coating material on top of the substrate.

Turn on the fan and spin coat the substrate for about 30 seconds time can vary depending on the substrate viscosity and coating thickness required.

Verify the coating by peeling off the PDMS membrane from the glass slide by tweezer left picture or analyze the microchannel profile by microscopy right panels.

In this tip a portable spin coater for microfluidic applications was developed using old electronic parts. A single fan can be re-used many times up to hundreds in our experience.

The amount of PDMS in form of droplets falling on the fan is quite limited. If necessary the fan can be cleaned after any use by simply rubbing it with a wipe soaked with some petroleum ether aka liquid paraffin or white petroleum.

In the worst cases very rarely occurring the fan can be easily replaced, since they are available for free by any old unused PC.

Hall, P. Underhill, and J. Halldorsson, E. Lucumi, R. Vecchione, G. Pitingolo, D. Guarnieri, A. 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.

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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.

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