Name: scalable 96-well plate based ipsc culture and production using a robotic liquid handling system
Uploaded: 2015-05-14T14:00:00
Duration: 8 min
Description: Watch this Scientific Journal Video about Scalable 96-well Plate Based iPSC Culture and Production Using a Robotic Liquid Handling System at JoVE.com

This work has been supported in part by NIH grants, R44 GM087784 and R01 HL109505. Authors thank the technical and OEM team at Gilson, INC for extended technical support.

The robotic liquid handling system, which for the following protocols was Gilson&#8217;s pipetmaX, facilitates automated, scalable stem cell culture and production while maintaining traditional adherent culture conditions in the 96-well plate format. Like traditional 6-well plate culture, iPSCs are grown with this protocol as a monolayer of distinct colonies but in the 96-well plate format. Each well can be isogenic or distinct and either configuration can be cultured in parallel since the robot can keep each well segregated. The typical robotic iPSC culture routine has two phases: a feeding program where cultures are fed on a customizable schedule and a passage program where the user chooses the dissociation method and splitting ratio thereby controlling seeding density and scale rate. The following protocols describe how a new culture is started, how feeding and passage are accomplished and the supplementary programs that facilitate iPSC culture.
Note: Please see the Materials List for specific recommendations on the matrix coating material and dissociation reagents validated for the following protocols.
1. Preparing Extracellular Matrix Gel Coated 96-well Plates
<ol>
	<li>Remove the packaging from six new, RT 96-well plates and place them on the liquid handling robot bed in positions 2, 3, 5, 7, 8, 9 (see Figure 1A for bed positions).</li>
	<li>Place a pre-chilled, 4 &#176;C 4-well trough in bed position 6.</li>
	<li>Load column 12 of the tip rack in bed position 1 with pre-chilled, 4 &#176;C 200 &#181;l pipet tips.</li>
	<li>Fill the first (left most) trough well in bed position 6 with 25 ml 4 &#176;C DMEM/F12 by pouring an aliquot using sterile technique. Alternatively, use a pipet to complete the transfer.</li>
	<li>Remove the 96-well plate lids and slide each into the specially designed plate lid holding rack (PLHR, see Figure 1A). Avoid contact with the inside of plate lids. 
	Note: Handling the inside of a plate lid or stacking the plate lids where the inside of one lid touches the outside of another is an excellent route for contamination.</li>
	<li>Using the robotic control touch pad, press the following series of buttons to start the well pre-wetting process:
	<ol>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;extracellular matrix gel Prewet&#8221; and tap next.</li>
		<li>User decision: Tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		Note: The robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
		<li>Tap &#8220;Run Protocol&#8221;.</li>
	</ol>
	</li>
	<li>During the pre-wetting process (step 1.6) continue to step 1.8.</li>
	<li>Thaw on ice one 4 mg aliquot of growth factor reduced extracellular matrix gel (GFRM) contained in a 50 ml conical tube stored at -80 &#176;C. Keep the aliquot on ice until ready to use. 
	Note: The GFRM will solidify if allowed to come to RT and will not be useful.</li>
	<li>Resuspend the 4 mg GFRM aliquot in 24 ml 4 &#176;C DMEM/F12 with a 25 ml glass pipet. When transferring the DMEM/F12, pause for 2-3 sec to allow the pipet to cool prior to resuspending the GFRM aliquot.</li>
	<li>When the pre-wetting process (step 1.6) is complete, continue to step 1.11.</li>
	<li>Transfer the 24 ml of DMEM/F12 GFRM mixture to the fourth well of the trough in bed position 6.</li>
	<li>Using the robotic control touch pad, press the following series of buttons to start the extracellular matrix gel coating process:
	<ol>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;extracellular matrix gel aliquot&#8221; and tap next.</li>
		<li>User decision: Tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		Note: The robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
		<li>Tap &#8220;Run Protocol&#8221;.</li>
	</ol>
	</li>
	<li>When step 1.12 is complete, replace the plate lids.</li>
	<li>Transfer the extracellular matrix gel coated 96-well plates to a 37 &#176;C, 5% CO2, humidified incubator and hold there for 24 hr at which point the plates will be ready to use or stored. 
	Note: Under extenuating circumstances, the newly coated extracellular matrix gel plates may be used after 15-30 min of incubation but O/N to 24 hr incubation is recommended.</li>
	<li>Repeat protocol steps 1.1, 1.3, 1.5-1.6, 1.10, and 1.12-14 until all of the DMEM/F12 with GFRM is used.</li>
</ol>
2. Storing Extracellular Matrix Gel Coated 96-well Plates
<ol>
	<li>Remove extracellular matrix gel coated 96-well plates from the 37 &#176;C, 5% CO2, humidified incubator.</li>
	<li>Optional: Allow the extracellular matrix gel coated 96-well plates to come to RT before proceeding to step 2.3. 
	Note: This optional step will reduce condensation build up when the plates are placed at 4 &#176;C.</li>
	<li>Place the plates on the liquid handling robot bed in positions 2, 3, 5, 7, 8, 9 (see Figure 1A for bed positions).</li>
	<li>Load column 12 of the tip rack in bed position 1 with RT&#160;200 &#181;l pipet tips.</li>
	<li>Place a four trough reservoir in bed position 6.</li>
	<li>Fill well 4 (rightmost) of the reservoir with RT DMEM/F12.</li>
	<li>Remove the 96-well plate lids and slide each into the PLHR.</li>
	<li>Using the robotic control touch pad, press the following combination of buttons:
	<ol>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;extracellular matrix gel aliquot&#8221; and tap next.</li>
		<li>User decision: Tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		Note: The robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
		<li>Tap &#8220;Run Protocol&#8221;.</li>
	</ol>
	</li>
	<li>Once complete, replace the plate lids and remove the plates from the machine bed.</li>
	<li>Wrap around the entire side of each extracellular matrix gel coated 96-well plate (where the lid meets the plate) with a one inch by 6 inch paraffin film strip. Be sure to stretch the paraffin film to create an airtight seal.</li>
	<li>Optional: Label the plates with the date of completed extracellular matrix gel coating, the extracellular matrix gel lot number, user name and any other information.</li>
	<li>Store the plates at 4 &#176;C where they will be good to use for up to two weeks. 
	Note: Plates have been successfully used beyond the two week limit, however, it is not recommended.</li>
</ol>
3. Performing a Stem Cell Colony Seeding Density Gradient in the 96-well Plate Format from a Live or Frozen Culture: How to Start or Restore Normal Passage Intervals to a Stem Cell Line
<ol>
	<li>If starting with a frozen culture, proceed to step 3.2. If starting with a live stem cell culture already on a plate, begin at step 3.5.</li>
	<li>Thaw the frozen culture by swirling the tube in a 37 &#176;C water bath until only a small chunk of ice remains.</li>
	<li>Optional: Dilute the thawed cells into 10 ml RT stem cell growth media (SCGM), centrifuge for 2 min at 300 x g, aspirate the supernatant media and resuspend the stem cell pellet in 7 ml fresh SCGM containing 10 &#181;M Y27632. 
	Note: This step eliminates carryover of the freezing media.</li>
	<li>Proceed to Step 3.6.</li>
	<li>If starting with live, already plated stem cells: passage the cells as normal using enzymatic or non-enzymatic dissociation. Try to harvest a total of 1.5-3 million cells; usually one well of a six-well plate. Centrifuge the harvested cells for 2 min at 300 x g, aspirate the supernatant media and resuspend the stem cell pellet in 7 ml fresh SCGM containing 10 &#181;M Y27632. Proceed to Step 3.6.</li>
	<li>Ensure the cells, either from a frozen or live culture, are suspended in 7 ml RT SCGM with 10 &#956;M Y27632.</li>
	<li>Load column 12 of the tip rack in bed position one with RT 200 &#181;l pipet tips.</li>
	<li>Place a four trough reservoir in bed position 2.</li>
	<li>Place one new extracellular matrix gel coated 96-well plate pre-warmed to 37 &#176;C in bed position 5 and remove the lid placing it in the PLHR.</li>
	<li>Transfer the 7 ml of resuspended cells to well 3 of the reservoir by pipet or sterile pour.
	<ol>
		<li>Using the robotic control touch pad, press the following combination of buttons:</li>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;Seeding Density Gradient&#8221; and tap next.</li>
		<li>User decision: Tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		Note: The robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
	</ol>
	</li>
	<li>Tap &#8220;Run Protocol&#8221;.</li>
	<li>When the dilution series is complete, re-cover the plate and place in a 37 &#176;C, 5% CO2, humidified incubator until the next feeding.</li>
	<li>Optional: Label the plate with the date, strain information, user name, media type and other pertinent information.</li>
	<li>Over the next several days, feed the 96-well plate as required using Protocol 4 below. 
	Note: A typical stem cell plate will require a full media change once every 24 hr and the SCGM does not contain Y27632 for feeding; only during the initial seeding after passage. Prior to feeding, check the plate for contamination and colony density. See step 3.16 for more on monitoring colony density.</li>
	<li>Over the next few days, several columns near the center of the plate (usually columns 5-8) will approach an ideal density for passage; to identify this density, compare the 96-well plate to the density range shown in Figure 3B with the following information in mind:
	<ol>
		<li>Passage when the space between the majority of the colonies is approximately 25% of adjacent colony diameter. 
		Note: The rationale to identify the time to passage is that another cycle of feeding and growth would result in colonies making contact, which should be avoided.</li>
	</ol>
	</li>
	<li>To start a uniformly diluted plate and begin regular culture, select a column that is at the ideal density and passage. See Protocol 5 to passage.</li>
</ol>
4. Feeding 96-well Plate Stem Cell Colonies
Note: The following protocol describes the feeding of one 96-well stem cell colony plate. The technique can be scaled up to accommodate 6 total plates in parallel.
<ol>
	<li>Remove the 96-well stem cell colony plate from the 37 &#176;C, 5% CO2, humidified incubator.</li>
	<li>Check the plate for contamination and cell density. In order to feed, ensure that the colony density is below the ideal passage density. See Figure 3B for information on density. 
	Note: Contamination may present as turbid media and be accompanied by an unpleasant smell. Small, obviously non-iPSC aggregates may also be visible under microscopic inspection. To evaluate cell density, see step 3.16.1 above.</li>
	<li>Place the 96-well stem cell colony plate in bed position 3 on a sloped, 37 &#176;C ramp.</li>
	<li>Load column 12 of the tip rack in bed position 1 with RT 200 &#181;l pipet tips.</li>
	<li>Place a 4-well trough in bed position 2.</li>
	<li>Fill well 3 of the trough in bed position 2 with 8 ml fresh, RT SCGM without Y27632 either by sterile pour or pipet transfer.</li>
	<li>Remove the 96-well stem cell colony plate lid placing it in the PLHR.</li>
	<li>Using the robotic control touch pad, press the following combination of buttons:
	<ol>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;Colony Feed One Plate&#8221; and tap next.</li>
		<li>User decision: tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		&#8203;Note, the robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
		<li>Tap &#8220;Run Protocol&#8221;.</li>
	</ol>
	</li>
	<li>When the feeding protocol is complete, re-cover the 96-well stem cell colony plate and return to the 37 &#176;C, 5% CO2, humidified incubator until the next feeding or passage is required. This is usually required after 24 hr. 
	Note: It is recommended to record the feeding event on the plate cover with the date, media type, user name and other pertinent information.</li>
</ol>
5. Passaging 96-well Plate Stem Cell Colonies
<ol>
	<li>After several feeding cycles the 96-well stem cell colony plate will be ready to passage. This protocol describes passage of one column to one 96-well plate, representing a 1:12 split ratio. The user can define the split ratio and select any number of wells or columns to split, including choosing wells that may not be adjacent.</li>
	<li>Compare the 96-well stem cell colony plate density to Figure 3B to determine whether to passage. The ideal passage density is when the space between the majority of colonies is approximately 25% of adjacent colony diameter or a subsequent feeding would result in the colonies growing into one another.</li>
	<li>Examine the 96-well stem cell colony plate under a microscope to ensure there is no contamination.</li>
	<li>Place the 96-well stem cell colony plate in bed position 3 on a sloped, 37 &#176;C ramp.</li>
	<li>Load columns 8-12 in the tip loading rack in bed position 1 with RT 200 &#181;l pipet tips.</li>
	<li>Place a 4-well trough in bed position 2.</li>
	<li>Leave trough well 1 empty, put 8.5 ml SCGM + 10 &#181;M Y27632 in well 2, put 3.5 ml 30% proteolytic and collagenolytic dissociation reagent (or EDTA based dissociation reagent) in well 3, put 4.5 ml PBS in well 4. 
	Note: All reagents can be at RT or 37 &#176;C. RT proteolytic and collagenolytic dissociation reagent at 30% diluted with PBS was used for the Adipose and Fibroblast derived iPSC culture described here.</li>
	<li>Place one new, pre-warmed 37 &#176;C extracellular matrix gel coated 96-well plate in bed position 5.</li>
	<li>Remove the 96-well stem cell colony plate lid, as well as the new 96-well plate lid and place them both in the PLHR.</li>
	<li>Using the robotic control touch pad, press the following combination of buttons:
	<ol>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;Colony Split 1:12 Column 1&#8221; and tap next.</li>
		<li>User decision: Tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		Note: The robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
		<li>Tap &#8220;Run Protocol&#8221;.</li>
	</ol>
	</li>
	<li>Once the passage protocol has finished, re-cover both plates.</li>
	<li>Optional: Label the new plate with the requisite information (i.e., date, cell line, passage number, media type, user name etc.).</li>
	<li>Return both plates to the 37 &#176;C, 5% CO2, humidified incubator until the next feeding or passage is required. Note: The next feeding is typically after 24 hr. 
	Note: Cells and colonies should attach to the plate within 2-3 hr of splitting and look very spread out. This is due to the presence of Rho-kinase inhibitor and the cells will condense and exhibit the characteristic stem cell morphology (i.e., cobblestone packed colonies with small cell volume to big nucleus ratio) after the first Rho-kinase inhibitor free feeding.</li>
</ol>
6. Harvesting 96-well Plate Stem Cells for Production
Note: Stem cell colonies may be harvested for use at any point during culture. Usually this occurs when the cells are ready to passage (see protocol 5). This protocol describes how to harvest eleven columns from a 96-well stem cell colony plate.
<ol>
	<li>Examine the 96-well stem cell colony plate under a microscope to ensure there is no contamination.</li>
	<li>Place the 96-well stem cell colony plate in bed position 3 on a sloped, 37 &#176;C ramp.</li>
	<li>Load columns 11 and 12 in the tip loading rack in bed position 1 with RT 200 &#181;l pipet tips.</li>
	<li>Place a 4-well trough in bed position 2.</li>
	<li>Leave trough well 1 empty, put 8.5 ml SCGM in well 2, put 3.5 ml 30% proteolytic and collagenolytic dissociation reagent (or EDTA based dissociation reagent) in well 3, put 4.5 ml PBS in well 4. 
	Note: All reagents can be at RT or 37 &#176;C.</li>
	<li>Remove the 96-well stem cell colony plate lid and place in the PLHR.</li>
	<li>Using the robotic control touch pad, press the following combination of buttons:
	<ol>
		<li>Tap &#8220;Run a Protocol&#8221;.</li>
		<li>Select &#8220;Colony Harvest Columns 2-12&#8221; and tap next.</li>
		<li>User decision: Tap &#8220;test run&#8221; to use the step-by-step wizard to pre-test the selected program. 
		Note: The robot does not run but rather the software tests the protocol for software problems. With established protocols, tapping, &#8220;skip setup&#8221; is acceptable.</li>
		<li>Tap &#8220;Run Protocol&#8221;. 
		Note: Once the collection procedure is complete, the cells will be in trough well 2 and ready for collection.</li>
	</ol>
	</li>
</ol>

The authors MKC, MJG, EEB, NJD and RO are or were at the time of development employees of InvivoSciences, INC which conceived of and developed the robotic protocols described here. TW is the CSO for InvivoSciences INC. SH is an employee of Gilson INC.

In the present study we developed a method of robotic culture for iPSC production that miniaturizes and automates feeding and passage in a 96-well plate format while also enabling efficient scale up. A liquid handling robot was used to routinely feed iPSC colonies and passage them at regular intervals by enzymatic dissociation. The robot was also programmed to produce extracellular matrix gel coated plates and perform a seeding density gradient. When fibroblast and adipose derived iPSCs were cultured in this system for greater than 20 passages, about 3 months, pluripotency was maintained, as were stable karyotypes. Both cell lines were capable of differentiation into cardiomyocytes. Together, the collection of protocols described here enables users with very little stem cell culture experience, or limited access to specialized culture equipment, to culture stem cells and achieve high cell production or multi-line handling with minimal labor and cost.
The robotic protocols detailed here provide a method for economical scale up of iPSC production and handling. One advantage is scalable stem cell culture in an established plate based format demonstrated here and previously44,47&#8211;49 to maintain pluripotency. While other formats of scalable stem cell culture yield pluripotent cells20,23,29 this plate based method requires essentially no change in culture format, like adaption to suspension, use of antifoaming chemicals or prolonged use of Rho-kinase inhibitor20. This plate based method can therefore rapidly, and predictably, accommodate new lines because the culture conditions are similar. As the use of iPSCs for disease modeling increases4,6,19,50, new iPSC lines are continuously generated. The techniques described here are best suited to culture new lines in medium to high volume and throughput because the barrier to transition onto the format is low. This is also true for the labor and equipment necessary to maintain the colonies. Finally, because the format and handling are similar to the environment used to derive and validate iPSC lines, culture performance, such as directed differentiation, is likely to be more predictable.
Figure 8 illustrates one strategy for scale up of stem cell production in 96-well plates. One plate (Cycle 0) is used to robotically seed 12 new plates, a 12 fold increase (Cycle 0 to 1). After several days of feeding, one of the twelve plates is passaged to seed another set of twelve plates (Cycle 1 to Cycle 2). The remaining eleven plates are now available for other uses and represent the production component of the system. At this rate, passage of one plate will provide 11 plates every cycle, or every 3-4 days. This methodology can be scaled more when multiple plates are passaged as shown in the transition from Cycle 2 to Cycle 3. In Cycle 3, two plates are always used to maintain the colony and seed 24 new plates. The remaining 22 plates are then available for use after every passage cycle. At any point in the maintenance cycle the user can seed more plates to increase production. One example would be to passage every column for two growth cycles. The first passage converts one plate into twelve plates, and each of the twelve plates is used to seed 144 plates. In this case, a user starts with one plate and after two passages, or about 1.5-2 weeks of culture, has 144 plates. For example, the fibroblast and adipose iPSC lines yield approximately 25-75 million cells per 96-well plate when ready to passage. 144 plates could therefore represent about 4-7 x 109 cells.
What is the labor burden for carrying 144 96-well plates robotically? One of the goals for this work was scalable stem cell production that reduced labor compared to traditional (i.e., 6-well plate) stem cell culture. The robot accomplishes this by alleviating the need to physically handle each plate during feeding and passaging. While 96-well plate production scales linearly like it would in traditional 6-well plates, the time a technician spends working with the plates is significantly less using robotic culture. The production efficiency of robotic culture is derived from this difference (Figure 9). It was found technicians could remove a 6-well plate from the incubator, check it on the microscope, then aspirate and feed the plate in approximately 3.5 minutes. In comparison, technicians spend approximately 30 sec&#160;removing a 96-well plate from the incubator and inspecting it on the microscope for density and contamination before placing it on the robot. With an expanded feeding protocol similar to the one described above, six 96-well plates can be loaded and fed, which takes approximately 21 min, or 3.5 min&#160;per plate. The total elapsed time to feed a plate is comparable between robotic and manual methods, but to hand feed 6 plates a technician is required for all 21 min&#160;whereas the technician spends only 3 min&#160;handling the six plates for the robot (a 30 sec&#160;inspection per plate). As Figure 9 shows, the time a technician spends handling 144 plates using the robot is about 1.2 hr as opposed to 8 hr manually and the robot will never make a mistake handling the plates or transferring liquids.
The 96-well iPSC culture methods described here provide a tractable platform for complex screening tasks. Because each well is genetically identical and seeded at the same density from the same initial pool, variables can be distributed across the plate and maintained by the robot with commercially available reservoirs to segregate conditions. This would be useful for experiments such as stem cell growth media development34,35,47, substrate or culture condition testing and toxicity studies utilizing stem cells51. Since each stem cell line is unique in terms of optimal colony size and passage requirements, one can use this system to modulate seeding density, feeding frequency and passage handling by manipulating the columns harvested, the mechanical agitation during passage and the feeding frequency. Iterating through these variables will allow the user to quickly find a set of conditions suitable for culture of a new line or develop new culture techniques. The robotic system is also capable of maintaining 96 parallel but independent stem cell colonies. When iPSC are derived from somatic cells or cloned after genetic manipulation, many parallel clones are screened to yield potential iPSC lines. Once single cells are seeded into 96-well plates, this system can feed and passage the 96-well plates keeping each colony segregated. This enables very high throughput when selecting potential clones and eliminates the difficulty in physically culturing 96 parallel lines. This method also allows scaled up production of each clone so that material is readily available for parallel analysis. Finally, we envision integrating these culture techniques with a robotic plate trafficking system that delivers plates to and from the incubator enabling fully automated culture. This could be accomplished with more complex robotics that allow for greater expansion since the basic protocols described here are transferable to other plate based systems.
The first critical steps to address in the protocols are ones that prevent and check for contamination such as steps 1.5 and 4.2. Contamination, as discussed above, can be successfully avoided if good sterile technique and common sense are utilized. It is imperative, regardless of stem cell culture format, that the operator check for contamination and doing so in this protocol at the indicated steps will significantly reduce the risk for contamination. Proper extracellular matrix gel application is a second essential step to the overall success of this protocol. Without properly coating the 96-well plates, the stem cells will not grow. One common issue is incomplete well coating. Experience has shown that air bubbles, electrostatic attraction and capillary action hinder complete well coating. The pre-wetting step (1.6) was developed to significantly improve well bottom coating and should not be skipped. This pre-wetting step appears to reduce the apparent hydrophobicity of the plastic 96-well plate such that when the extracellular matrix gel coating is applied it is evenly distributed across the well bottom and sides. It is also recommended to gently tap the 96-well plates against a clean hand once coated to ensure even extracellular matrix gel solution distribution. The dissociation parameters are also important to optimize. Extended incubation with enzyme or EDTA based dissociation reagent will result in single cells which may or may not be the goal during passage. Therefore, the operator should pay special attention to how dissociation time, trituration force, and wash repetitions affect colony morphology and health and adjust accordingly.
Understanding the limitations to the techniques described here are paramount to successful operation. As in any other cell culture setting, the use of 96-well plates presents a relatively high contamination risk. As described above, a technician is handling each plate during feeding and passaging; for example, when opening the lid, putting the plate on the robot bed, and checking the plate on the microscope. Therefore, as noted after step 1.5, it is imperative not to touch the inside of any plate lid or expose this part to any potentially contaminated surface such as the sloped heating blocks or vertical pins on the bed. During protocol development this limitation was discovered and was the impetus to develop a rack to hold plate lids to maintain sterility. Likewise, the trough reservoirs, pipet tips and other supplies on the liquid handling robot bed are potential sources of contamination because they are open to environment and handled by the technician. Therefore, good sterile technique should be applied such as reducing movements over exposed culture materials or open liquids. Another limitation is that when the protocol is scaled up, visually checking every well of &#62;100 96-well plates is cumbersome. This can be dealt with by visually inspecting the whole plate for signs of contamination, such as turbid media, to identify suspicious wells. For future scale up, use of an automated microscope and indicator of cell growth and potential contamination will be incorporated.
In summary, the high-throughput 96-well based platform described here offers a reproducible, high fidelity method for stem cell culture and production in a plate format. This method reduces the requisite experience, equipment necessary and labor time dedicated to stem cell culture while maintaining the benefits of traditional adherent culture.

The use of human induced pluripotent stem cells (iPSCs) has increased significantly since their derivation in 20071 for compound testing, regenerative medicine and disease modeling2&#8211;6. This demand comes from iPSC&#8217;s capacity to yield large numbers of pluripotent cells that can be differentiated into somatic cells at an unparalleled quantity. As directed differentiation techniques improve7&#8211;10 and development of human cell or tissue modeling and cell therapy increases11&#8211;14, so does the requirement for mass-production of high quality iPSCs. It is widely cited that among other maladies, myocardial infarction or b-cell functional replacement will require hundreds of millions to billions of iPSC derived somatic cells15&#8211;18. Furthermore, increasingly complex 3D tissue modeling for drug discovery and therapy will demand large numbers of cells9,13,19. In all of these examples, defined, uniform and reproducible iPSCs must be available and straightforward to produce.
In order to produce stem cell derived biopharmaceutics and cells for tissue engineering and transplantation, a cost-effective cell-manufacturing technology is essential. Scaled production of iPSCs has focused on suspension culture20&#8211;25 or suspension with the use of microcarrier substrates26&#8211;28 partly because these techniques are successfully deployed for large scale, non-iPSC, eukaryotic based production. Several groups have demonstrated suspension culture systems that yield pluripotent stem cells20,21,23,24,29. However, these approaches utilize expensive and complex systems not readily available to researchers developing novel differentiation programs, driving tissue engineering and doing laboratory scale research. Furthermore, suspension and microcarrier iPSC cultures require adaptation and techniques or chemicals not found in traditional adherent iPSC culture such as continuous use of Rho-kinase inhibitor, antifoaming agents and filtration to regulate aggregation. Physical stress to the cells is more prevalent in suspension culture, introduced by mechanical agitation as well as during microcarrier collision. These issues limit the predictability and speed at which newly generated stem cell lines can be cultured in suspension. Others have developed robotic plate handling systems that mimic plate culture techniques30,31 but these platforms demand significant investments for equipment and expertise to operate in addition to the fundamental challenges associated with stem cell culture.
The following work describes the development and practice of a scalable iPSC culture system that utilizes a self-contained, standard 8-channel robotic liquid handler and 96-well plates. This method was designed to bridge laboratory scale iPSC culture and high-volume iPSC production (e.g., 107 &#8211; 1.5 x 109 cells per week per technician) enabling those developing new iPSC technologies to easily scale up production without large hardware or labor investments. This method is relatively inexpensive to set up, requires little to no stem cell culture, programming or engineering experience, has a small equipment footprint without the need for a dedicated cell culture hood, and utilizes standard cell culture equipment to enable medium to high throughput automated cell production. The aim was to develop a system capable of scalable stem cell culture that can be utilized by laboratories new to stem cell culture, those who find manual cultivation a barrier to developing their ideas or those wanting an economical means to produce large numbers of stem cells. The platform presented here removes technician influence on stem cell culture and normalizes the feeding and passage procedures to enable consistent stem cell production.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>96-well plates</td>
			<td>Corning</td>
			<td>3596</td>
			<td>96-well; Well volume: 360uL; Cell growth area: 0.32cm2; Individually wrapped</td>
		</tr>
		<tr>
			<td>Seahorse Trough</td>
			<td>SeahorseBio</td>
			<td>201308-100</td>
			<td>Reservoir 4 Clear Part Poly Proplene 73Ml 25/Cs</td>
		</tr>
		<tr>
			<td>Gilson Tips</td>
			<td>Gilson</td>
			<td>F167023</td>
			<td>10 racks gilson 96tips D200 tips</td>
		</tr>
		<tr>
			<td>DMEM/F12</td>
			<td>Life Technologies</td>
			<td>12500062</td>
			<td>DMEM/F12 powder.&#160; Resuspend in 1 L purified, cell culture grade water and sterile filter.</td>
		</tr>
		<tr>
			<td>Growth Factor Reduced Matrigel</td>
			<td>Corning</td>
			<td>354230</td>
			<td>Referred to as, &#34;extracellular matrix gel&#34; in the text.&#160; Matrigel GFR, 10 mL</td>
		</tr>
		<tr>
			<td>mTeSR1</td>
			<td>StemCell Technologies</td>
			<td>5857</td>
			<td>mTeSR1 Complete Kit for hES Maintenance.</td>
		</tr>
		<tr>
			<td>E8 Media</td>
			<td>StemCell Technologies</td>
			<td>5940</td>
			<td>TeSR-E8 Kit for hESC/hiPSC Maintenance</td>
		</tr>
		<tr>
			<td>Y27632</td>
			<td>AdooQ BioScience</td>
			<td>A11001-50</td>
			<td>Rock inhinitor Y-27632 2HCI</td>
		</tr>
		<tr>
			<td>Accutase</td>
			<td>Innovative Cell Technologies</td>
			<td>ACCUTASE</td>
			<td>Referred to as, &#34;proteolytic and collagenolytic dissociation reagent&#34; in the text.&#160; Accutase 500 mL sterile cell solution</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Fisher</td>
			<td>SH30256FS</td>
			<td>PBS w/o CA MG 500 mL, 6/pk</td>
		</tr>
		<tr>
			<td>Gilson PIPETMAX</td>
			<td>Gilson</td>
			<td>PIPETMAX</td>
			<td>http://www.gilson.com/en/AI/Products/13.290/Default.aspx#.VCGwRBZmYSk</td>
		</tr>
	</tbody>
</table>

scalable 96-well plate based ipsc culture and production using a robotic liquid handling system

Continued advancement in pluripotent stem cell culture is closing the gap between bench and bedside for using these cells in regenerative medicine, drug discovery and safety testing. In order to produce stem cell derived biopharmaceutics and cells for tissue engineering and transplantation, a cost-effective cell-manufacturing technology is essential. Maintenance of pluripotency and stable performance of cells in downstream applications (e.g., cell differentiation) over time is paramount to large scale cell production. Yet that can be difficult to achieve especially if cells are cultured manually where the operator can introduce significant variability as well as be prohibitively expensive to scale-up. To enable high-throughput, large-scale stem cell production and remove operator influence novel stem cell culture protocols using a bench-top multi-channel liquid handling robot were developed that require minimal technician involvement or experience. With these protocols human induced pluripotent stem cells (iPSCs) were cultured in feeder-free conditions directly from a frozen stock and maintained in 96-well plates. Depending on cell line and desired scale-up rate, the operator can easily determine when to passage based on a series of images showing the optimal colony densities for splitting. Then the necessary reagents are prepared to perform a colony split to new plates without a centrifugation step. After 20 passages (~3 months), two iPSC lines maintained stable karyotypes, expressed stem cell markers, and differentiated into cardiomyocytes with high efficiency. The system can perform subsequent high-throughput screening of new differentiation protocols or genetic manipulation designed for 96-well plates. This technology will reduce the labor and technical burden to produce large numbers of identical stem cells for a myriad of applications.

Watch this Scientific Journal Video about Scalable 96-well Plate Based iPSC Culture and Production Using a Robotic Liquid Handling System at JoVE.com

Scalable 96-well Plate Based iPSC Culture and Production Using a Robotic Liquid Handling System

The protocols described allow laboratories to perform scalable, adherent stem cell culture in high throughput with minimal labor, experience and equipment investment cost using a programmable liquid handling robot and 96-well plates. iPSCs passaged more than 20 times on this system maintained pluripotency, normal karyotypes and differentiated into cardiomyocytes.

Continued advancement in pluripotent stem cell culture is closing the gap between bench and bedside for using these cells in regenerative medicine, drug ...

scalable-96-well-plate-based-ipsc-culture-production-using-robotic

Research

JoVE Journal

Developmental Biology

Alginate Encapsulation of Pluripotent Stem Cells Using a Co-axial Nozzle

Pluripotent stem cells (PS cells) are the focus of intense research due to their role in regenerative medicine and drug screening. However, the development of a mass culture system would be required for using PS cells in these applications. Suspension culture is one promising culture method for the mass production of PS cells, although some issues such as controlling aggregation and limiting shear stress from the culture medium are still unsolved. In order to solve these problems, we developed a method of calcium alginate (Alg-Ca) encapsulation using a co-axial nozzle. This method can control the size of the capsules easily by co-flowing N2 gas. The controllable capsule diameter must be larger than 500 &#181;m because too high a flow rate of N2 gas causes the breakdown of droplets and thus heterogeneous-sized capsules. Moreover, a low concentration of Alg-Na and CaCl2 causes non-spherical capsules. Although an Alg-Ca capsule without a coating of Alg-PLL easily dissolves enabling the collection of cells, they can also potentially leak out from capsules lacking an Alg-PLL coating. Indeed, an alginate-PLL coating can prevent cellular leakage but is also hard to break. This technology can be used to research the stem cell niche as well as the mass production of PS cells because encapsulation can modify the micro-environment surrounding cells including the extracellular matrix and the concentration of secreted factors.

We established a method of encapsulating pluripotent stem cells (PS cells) into alginate hydrogel capsules using a co-axial nozzle. This prevents cells from aggregating excessively and limits the shear stress experienced by cells in suspension culture. The technique is applicable to the mass production of PS cells as well as research on stem cell niche.

Pluripotent stem cells (PS cells) are the focus of intense research due to their role in regenerative medicine and drug screening.  However, the ...

Method of Studying Palatal Fusion using Static Organ Culture

Cleft lip and palate are among the most common of all birth defects. The secondary palate forms from mesenchymal shelves covered with epithelium that adheres to form the midline epithelial seam (MES). The theories suggest that MES cells follow an epithelial to mesenchymal transition (EMT), apoptosis and migration, making a fused palate 1. Complete disintegration of the MES is the final essential phase of palatal confluence with surrounding mesenchymal cells. We provide a method for palate organ culture. The developed in vitro protocol allows the study of the biological and molecular processes during fusion. The applications of this technique are numerous, including evaluating responses to exogenous chemical agents, effects of regulatory and growth factors and specific proteins. Palatal organ culture has a number of advantages including manipulation at different stages of development that is not possible using in vivo studies.

Studies of palate development are motivated by the incidence of cleft palate, a birth defect that imposes a tremendous health burden and can leave lasting disfigurement. We demonstrate here a technique to culture palatal shelves that can be used to study different signaling pathways involved in palatal development and fusion.

Cleft lip and palate are among the most common of all birth defects. The secondary palate forms from mesenchymal shelves covered with epithelium that ...

Functional Cloning Using a Xenopus Oocyte Expression System

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in abundance, may not be secreted but tethered to the signaling cell(s), or may require the presence of binding partners or upstream regulatory molecules. Thus in a search for gene products capable of eliciting an early lens-inductive response in competent ectoderm, we utilized an expression cloning system that would allow identification of paracrine or juxtacrine factors as well as transcriptional or other regulatory proteins. Pools of mRNA were injected into Xenopus oocytes, and responding tissue placed directly on the oocytes and co-cultured. Following functional cloning of ldb1 from a neural plate stage cDNA library based on its ability to elicit the expression of the early lens placode marker foxe3 in lens-competent animal cap ectoderm, we characterized the mRNA expression pattern, and assayed developmental progression following overexpression or knockdown of ldb1. This system is suitable in a very wide variety of contexts where identification of an inducer or its upstream regulatory molecules is sought using a functional response in competent tissue.

Functional Cloning Using a Xenopus Oocyte Expression System

We describe a Xenopus oocyte and animal cap system for the expression cloning of genes capable of inducing a response in competent ectoderm, and discuss techniques for the subsequent analysis of such genes. This system is useful in the functional identification of a wide range of gene products.

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

Large-Scale Production of Cardiomyocytes from Human Pluripotent Stem Cells Using a Highly Reproducible Small Molecule-Based Differentiation Protocol

Maximizing the benefit of human pluripotent stem cells (hPSCs) for research, disease modeling, pharmaceutical and clinical applications requires robust methods for the large-scale production of functional cell types, including cardiomyocytes. Here we demonstrate that the temporal manipulation of WNT, TGF-&#946;, and SHH signaling pathways leads to highly efficient cardiomyocyte differentiation of single-cell passaged hPSC lines in both static suspension and stirred suspension bioreactor systems. Employing this strategy resulted in ~ 100% beating spheroids, consistently containing &#62; 80% cardiac troponin T-positive cells after 15 days of culture, validated in multiple hPSC lines. We also report on a variation of this protocol for use with cell lines not currently adapted to single-cell passaging, the success of which has been verified in 42 hPSC lines. Cardiomyocytes generated using these protocols express lineage-specific markers and show expected electrophysiological functionalities. Our protocol presents a simple, efficient and robust platform for the large-scale production of human cardiomyocytes.

Here, we present a robust, fast and scalable cardiomyocyte differentiation protocol for human pluripotent stem cells (hPSCs). Cardiomyocytes derived using this large-scale method can provide sufficient cell numbers for their effective use in human cardiovascular disease modeling, high-throughput drug screening, and potentially clinical applications.

Maximizing the benefit of human pluripotent stem cells (hPSCs) for research, disease modeling, pharmaceutical and clinical applications requires robust ...

The Production of Pluripotent Stem Cells from Mouse Amniotic Fluid Cells Using a Transposon System

Induced pluripotent stem (iPS) cells are generated from mouse and human somatic cells by forced expression of defined transcription factors using different methods. Here, we produced iPS cells from mouse amniotic fluid cells, using a non-viral-based transposon system. All obtained iPS cell lines exhibited characteristics of pluripotent cells, including the ability to differentiate toward derivatives of all three germ layers in vitro and in vivo. This strategy opens up the possibility of using cells from diseased fetuses to develop new therapies for birth defects.

In this study, we generate induced pluripotent stem cells from mouse amniotic fluid cells, using a non-viral-based transposon system.

Induced pluripotent stem (iPS) cells are generated from mouse and human somatic cells by forced expression of defined transcription factors using ...

A Device for Performing Cell Migration/Wound Healing in a 96-Well Plate

The cell migration/wounding assay is a commonly used method to study cell migration and other biological processes, such as angiogenesis and tumor metastasis. In this assay, cells are grown to form a confluent monolayer and a mechanical wound is created by scratching with a device. Then the migration rate of the cells towards the denuded area can be monitored by imaging. Our 8-channel mechanical wounder is designed to tackle most of the problems associated with the cell migration assay. Firstly, our wounder can be easily sterilized by autoclaving or with common disinfectants. Secondly, the individual adjustable pins allow even contact with the cell culture plate so that sharp and reproducible wounds can be created. Thirdly, the guiding bars on both sides of the wounder ensure consistent wounding position in each well. The use of disposable plastic pipette tips for wounding can further provide better handling of the wounder as well as to minimize cross-contamination. In conclusion, our cell wounder can provide researchers with a user friendly and reproducible device for performing the cell migration assay using the standard 96-well culture plate.

Here, we present a protocol to perform a semi-high throughput cell migration assay on a 96-well cell culture plate. This protocol is a fast, simple and economical method to create consistent scratch wounds on a cell monolayer.

The cell migration/wounding assay is a commonly used method to study cell migration and other biological processes, such as angiogenesis and tumor ...

Defined Xeno-free and Feeder-free Culture Conditions for the Generation of Human iPSC-derived Retinal Cell Models

The production of specialized cells from pluripotent stem cells provides a powerful tool to develop new approaches for regenerative medicine. The use of human-induced pluripotent stem cells (iPSCs) is particularly attractive for neurodegenerative disease studies, including retinal dystrophies, where iPSC-derived retinal cell models mark a major step forward to understand and fight blindness. In this paper, we describe a simple and scalable protocol to generate, mature, and cryopreserve retinal organoids. Based on medium changing, the main advantage of this method is to avoid multiple and time-consuming steps commonly required in a guided differentiation of iPSCs. Mimicking the early phases of retinal development by successive changes of defined media on adherent human iPSC cultures, this protocol allows the simultaneous generation of self-forming neuroretinal structures and retinal pigmented epithelial (RPE) cells in a reproducible and efficient manner in 4 weeks. These structures containing retinal progenitor cells (RPCs) can be easily isolated for further maturation in a floating culture condition enabling the differentiation of RPCs into the seven retinal cell types present in the adult human retina. Additionally, we describe quick methods for the cryopreservation of retinal organoids and RPE cells for long-term storage. Combined together, the methods described here will be useful to produce and bank human iPSC-derived retinal cells or tissues for both basic and clinical research.

The production of specialized retinal cells from pluripotent stem cells is a turning point in the development of stem cell-based therapy for retinal diseases. The present paper describes a simple method for an efficient generation of retinal organoids and retinal pigmented epithelium for basic, translational, and clinical research.

The production of specialized cells from pluripotent stem cells provides a powerful tool to develop new approaches for regenerative medicine. The use of ...

An In Vitro Model of a Parallel-Plate Perfusion System to Study Bacterial Adherence to Graft Tissues

Various valved conduits and stent-mounted valves are used for right ventricular outflow tract (RVOT) valve replacement in patients with congenital heart disease. When using prosthetic materials however, these grafts are susceptible to bacterial infections and various host responses.
Identification of bacterial and host factors that play a vital role in endovascular adherence of microorganisms is of importance to better understand the pathophysiology of the onset of infections such as infective endocarditis (IE) and to develop preventive strategies. Therefore, the development of competent models to investigate bacterial adhesion under physiological shear conditions is necessary. Here, we describe the use of a newly designed in vitro perfusion chamber based on parallel plates that allows the study of bacterial adherence to different components of graft tissues such as exposed extracellular matrix, endothelial cells and inert areas. This method combined with colony-forming unit (CFU) counting is adequate to evaluate the propensity of graft materials towards bacterial adhesion under flow. Further on, the flow chamber system might be used to investigate the role of blood components in bacterial adhesion under shear conditions. We demonstrated that the source of tissue, their surface morphology and bacterial species specificity are not the major determining factors in bacterial adherence to graft tissues by using our in-house designed in vitro perfusion model.

An In Vitro Model of a Parallel-Plate Perfusion System to Study Bacterial Adherence to Graft Tissues

We describe an in-house designed in vitro flow chamber model, which allows the investigation of bacterial adherence to graft tissues.

Various valved conduits and stent-mounted valves are used for right ventricular outflow tract (RVOT) valve replacement in patients with congenital heart ...

Three-dimensional Angiogenesis Assay System using Co-culture Spheroids Formed by Endothelial Colony Forming Cells and Mesenchymal Stem Cells

Studies in the field of angiogenesis have been aggressively growing in the last few decades with the recognition that angiogenesis is a hallmark of more than 50 different pathological conditions, such as rheumatoid arthritis, oculopathy, cardiovascular diseases, and tumor metastasis. During angiogenesis drug development, it is crucial to use in vitro assay systems with appropriate cell types and proper conditions to reflect the physiologic angiogenesis process. To overcome limitations of current in vitro angiogenesis assay systems using mainly endothelial cells, we developed a 3-dimensional (3D) co-culture spheroid sprouting assay system. Co-culture spheroids were produced by two human vascular cell precursors, endothelial colony forming cells (ECFCs) and mesenchymal stem cells (MSCs) with a ratio of 5 to 1. ECFCs+MSCs spheroids were embedded into type I collagen matrix to mimic the in vivo extracellular environment. A real-time cell recorder was utilized to continuously observe the progression of angiogenic sprouting from spheroids for 24 h. Live cell fluorescent labeling technique was also applied to tract the localization of each cell type during sprout formation. Angiogenic potential was quantified by counting the number of sprouts and measuring the cumulative length of sprouts generated from the individual spheroids. Five randomly-selected spheroids were analyzed per experimental group. Comparison experiments demonstrated that ECFCs+MSCs spheroids showed greater sprout number and cumulative sprout length compared with ECFCs-only spheroids. Bevacizumab, an FDA-approved angiogenesis inhibitor, was tested with the newly-developed co-culture spheroid assay system to verify its potential to screen anti-angiogenic drugs. The IC50 value for ECFCs+MSCs spheroids compared to the ECFCs-only spheroids was closer to the effective plasma concentration of bevacizumab obtained from the xenograft tumor mouse model. The present study suggests that the 3D ECFCs+MSCs spheroid angiogenesis assay system is relevant to physiologic angiogenesis, and can predict an effective plasma concentration in advance of animal experiments.

Three-dimensional co-culture spheroid angiogenesis assay system is designed to mimic the physiologic angiogenesis. Co-culture spheroids are formed by two human vascular cell precursors, ECFCs and MSCs, and embedded in collagen gel. The new system is effective for evaluating angiogenic modulators, and provides more relevant information to the in vivo study.

Studies in the field of angiogenesis have been aggressively growing in the last few decades with the recognition that angiogenesis is a hallmark of more ...