Development of a Hydroponics System toImpose Osmotic Stress on Arabidopsis Plant

Mark Slater & Mike Gosney

Research under Michael Mickelbart, Purdue University Plant Science

Introduction My research this semester focused on optimizing the design of a hydroponics system in order to run tests on Arabidopsis thaliana. The genotype of A. thaliana we used was Columbia-0 (Col-0). The goal is to use PEG 6000 in the hydroponic nutrient solutions in order to induce drought stress and from that to gather drought response data. Hydroponics is a method of growing plants without using soil as a medium and instead growing the plants in a mineral nutrient solution. This can be done with or without a medium. Drought stress was shown to inhibit plant photosynthesis in A. thaliana, which inhibits in two ways: stomatal and non-stomatal (He et al, 2014). It causes stomatal closure which limits gas exchange. Plants require carbon dioxide as a reactant in the photosynthetic reaction, so limiting the gas exchange limits how much photosynthesis the plant can undergo. PEG 6000 is polyethylene glycol, which is a water soluble polymer with a molecular weight of 6000 g/mol. It is used as an inert, non-ionic solute to study the relationship between water and plants. We will use PEG with a high molecular weight (6000) because it cannot penetrate into the root apoplasts whereas the lower molecular weights of PEG can (Carpita et al, 1979). In order to use PEG 6000 to run tests, we dissolved different concentrations into the nutrient solutions that the A. thaliana plants were growing in. Adding PEG to a solution increases its osmolality, which are the number of dissolved particles in solution. Higher PEG concentrations result in increased osmolality, and increasing the osmolality of a solution increases the osmotic pressure (OP) of the solution. Osmotic pressure is a colligative property of the nutrient solution and is dependent on the amount of dissolved solutes. The total amount of ions of dissolved salts in the nutrient solution exert a force, which is OP. To measure osmolality, we used a vapor pressure osmometer. This type of osmometer works by plotting concentration vs electrical differential from standard solution concentrations and deriving the osmolality of the unknown solution from the plot. Another osmotic solute that is used in research is sucrose. It generates a lower OP than PEG of similar concentration. This suggests that the effects of PEG on OP reflects specific characteristics of the polymer (Money, 1989).

Expt 1 Rockwool Medium and PEG Buckets Objectives

The objective of this experiment is to create a functioning hydroponics system and address any issues that arise to optimize the design. We are testing whether we can improve upon the system design as well as other methods, such as medium used.

Materials and methodsThis system was broken into two parts: the germination system and the mature growth

system. For this experiment, the seeds were grown in rockwool medium inside 1.5 mL centrifuge tubes. Each tube had its cap cut off and we also clipped the bottom of each tube at the 0 mL mark (Figure 1). I then filled each tube with rockwool (Figure 1). Since rockwool is naturally high in salts, it required soaking before use. After constructing the seed bed, I placed each germination tube in it and filled with about 5.5 L of DI water (Figure 2). I then let it sit for a few days to soak and to draw out the salts from the rockwool. To put together the seed bed, I cut out foam board that was larger than the perimeter of the 5.5 L reservoir in order to rest on top of it. I then cut out many holes for the 1.5 mL centrifuge tube to go in the foam to be suspended by their caps. After letting them soak for a few days I placed A. thaliana seeds in each germination tube and covered them with plastic wrap to create a moist environment (Figure 3). To do this, seeds that had been stratified for no less than 3 days and were sown onto the top of the rockwool using toothpicks. After that, we connected the reservoir’s attached tubing to an air pump. The seeded tubes were placed in holes drilled into a sheet of foam insulation about .6 inches thick that sits on top of a 5.5 L plastic container filled with nutrient solution. Each piece of foam board is about 18 inches by 13 inches. This method did not result in very successful germination, but I had enough surviving plants to move them into the larger system.

There were 42 surviving plants that we could move to the larger system. We were testing 6 PEG concentrations, so we used 6 1.5 L buckets, each holding 7 plants. The buckets had PEG concentrations of 0%, 5%, 10%, 15%, 20%, and 25%. In order to make solutions with these different concentrations of PEG, we dissolved the weights listed in Table 2 of PEG into the bottles of nutrient solution. Each bucket contained equals amount of mixed nutrient solution made from Stock Solutions 1-6 listed in Table 1. In order to fully dissolve the higher concentrations of PEG, I heated and shook the bottles of solution. I wrapped each bucket in brown paper to reduce algal growth (Figure 4). I cut more foam board to fit inside the buckets and they floated on the water. I also covered each with aluminum to further reduce algal growth. To complete the system, I placed 7 plants in each bucket and connected each of the buckets’ tubing to an air pump (Figure 5). All systems were set up in growth chamber 3 in the teaching growth chamber in Horticulture Greenhouse. The light cycle was set up for 7 am to 5 pm and the temperature was 21 degrees Celsius. I had access to both sides of chamber 3 as well as both shelves in the unit. Data was collected on each bucket periodically and the pH was raised or lowered as necessary to get in the 5.8-6.0 range, which is the optimal range for nutrient uptake.

Figures

Table 1. Stock Nutrient Solutions

Table 2. PEG 6000 solutions

Figure 1. Rockwool Filled Germination Tube.

Figure 2. Soaking rockwool medium

Figure 3. Assembled germination system

Figure 4. Bucket system exterior

Figure 5. Completed bucket system

Results

Figure 6. EC Data for Bucket System

Figure 7. pH Data for Bucket System

Figure 8. Dry Weights of Bucket System

After a few days, many of the seeds in each bucket with a 10% concentration or higher began to die off. After 2 weeks, all of the plants in 15% PEG and higher were dead. The control bucket with 0% PEG was still thriving after 2 weeks, the 5% bucket only had a few die, and the 10% bucket had only one surviving plant. After 3 weeks, I dismantled this system in order to collect data from the plants. The dry weights of each surviving plant are shown in Figure 8. Clearly any concentration of PEG showed reduced growth of the plants when compared to the control.

As shown in Figure 6, the Electrical Conductivity (EC) for the buckets generally increased as time went on, with the 0% bucket having the largest increases. This happens even though the pH is maintained in the 5.8-6.0 range as shown in Figure 7. This is likely due to the fact that the buckets were not refilled as the plants took in water. So the data gathered on later dates had less water but still the same amount of PEG in each bucket.

Problems/potential solutionsThe first problem we encountered was the fact that so few seeds successfully germinated.

We believed that this was caused by the high amounts of salts in rockwool and we did not soak them thoroughly enough. If we were to use rockwool again we would soak them for longer than a couple days and switch out the water we were using for soaking a couple times throughout. Another issue we saw was that there was no easy way to transfer the seedlings into the larger system, and as a result, some of the roots were damaged in the transfer process. This is the most likely cause of non-uniformity between plants in the same PEG concentration bucket. Using a different way of germination to allow easier transfer was the potential solution we looked into. The buckets also had no way of keeping the roots separated, and it ended with a tangled mess of

roots below the bed. This would make it more difficult to sample roots. Some of the buckets had significant algal growth which also negatively affected the growth of the plants.

Expt 2 Agar Germination Caps and Larger System

ObjectivesThe objective of this experiment is to test a new system for germination and whether we

should adopt it or not. If we decide to use this new system, we will be looking for ways to improve it.

Materials and methodsWe decided to use a different system in order to have more success. Conn et al. (2013)

found a good design of a hydroponics system for A. thaliana, and we based our system off theirs. To construct this system, we had to create our own medium. We used plant agar (.7% weight/volume) dissolved in nutrient solution in a 100 mL container to create the agar medium. I used Stock Solutions 1-6 (Table 1) to create 100 mL of solution with dissolved agar and filled the rest up with DI water to the 100 mL mark. Often it required about 1 minute of time in the microwave, in 10 second intervals to prevent boiling over, to fully dissolve the agar in the solution. This solution will re solidify after a while and can be stored in the refrigerator for a month. To re use, simply loosen the cap and microwave it for about 1 minute, again at about 10 second intervals. This should return it to its liquid state and be suitable for use.

This method called for using the caps of the 1.5 mL centrifuge tubes as the seed container. To create the germination caps, I cut off the caps to each tube, keeping the lid connector intact. With this intact, it allows for easy pick up for the cap with tweezers. After that, I punched a hole in the center of the flat part of each cap. I then placed each cap flat side down on clear tape and pressed them down to create a seal (Figure 9) . In these, I placed ~300 microliters of the germination agar using a pipette and let solidify. You want to overfill each germination cap to form a little bubble on top. This bubble will help keep the agar moist which is essential for supplying nutrients to the seedling (Figure 10). If there is not enough agar in the cap or some spills off, you can add more to it. After the agar solidifies in the caps, about 10 minutes, you can remove them from the clear tape and place into the holder. We used a small plastic tray that holds 50 caps that fit over a rectangular reservoir (Figure 11). We then placed 3 seeds in the agar on the punched out hole on the germination caps. We used 3 to ensure at least one of the seeds germinated. We then filled the reservoir with enough water to submerge the agar bubbles, which was ~650 mL. Each plastic reservoir and lid hold 50 seeds. The caps float on water as well so you don't need to worry about the plants being flooded. We then covered the trays with plastic wrap (Figure 12). The seedlings should grow for about 14 days before puncturing the plastic wrap. At 21 days, they should be large enough to move to the larger system. I germinated at least one seed bed per week in order to have an abundance of seedlings at different stages in their growth cycles to test.

To construct the larger system, we used the same 5.5 L reservoirs with tubing as in Experiment 1. They were covered with a hard plastic sheet that had holes drilled in it that fit 50 mL centrifuge tubes, but were small enough that their caps could not fall through. This allowed the 50 mL centrifuge tubes to be suspended in the nutrient solution by the plastic (Figure 14). We used the 50 mL centrifuge tubes and their lids to hold the smaller germination caps. The caps had been cut with a 3/8 inch drill bit and smoothed down which snugly fit the smaller caps with the seedling and agar bubble (Figure 13). We then screwed them back onto the 50 mL tubes, which had holes punched throughout them to allow water and oxygen to get to the roots. At the time of transferring, the roots should have penetrated the agar bubble. We suspended each tube in the full concentration nutrient solution. We covered the plastic sheet with aluminum foil to prevent algal growth. After constructing everything, I connected the tubing in the reservoir to the air pump.

Figures

Figure 9. Germination Cap Setup

Figure 10. Filled Germination Caps with Bubbles

Figure 11. Germination Caps in Plastic Reservoir

Figure 12. Completed Germination Bed

Figure 13. 50 mL Centrifuge Tubes and Caps

Figure 14. Design of Larger System

ResultsAfter about a week the seedlings were getting too large to have 3 in each cap so I thinned

them to one seed per cap by removing the ungerminated seeds or weaker seedlings. To do this, I just used tweezers to remove the weaker plants or seeds. The first batch of 50 seeds using this new system nearly all died following the thinning phase. Because of this we did not have any plants that survived long enough to test the larger system.

Problems/potential solutionsWe thought that the thinning process was too stressful on the young plants and so we

decided to test it.

Expt 3 Thinning and Optimal Germination Nutrient Solution Concentration

ObjectivesThe first objective of this experiment is to determine whether or not the thinning phase

was killing the plants. To do this, we will have a control bed of seeds that we will not thin and allow to grow unchecked. For future plantings we will also only use 1 seed per cap to make thinning unnecessary. The second objective of this experiment is to determine the optimal concentration of nutrient solution for germinating A. thaliana. We will test full concentration nutrient solution and 50% diluted solution.

Materials and methodsThis experiment followed the same methods as the Experiment 2, with a few changes.

We created new seed beds to test whether the thinning phase was causing the death of the plants. We also created new ones to test the optimal concentration of nutrient solution while germinating. The control bed used had 3 seeds in it and did not undergo the thinning process. In the other beds, I used 1 seed per cap to prevent the necessity of thinning. If the control bed continued to grow past when the previous beds died, then it would indicate that the thinning was in fact killing them. I also prepared other seed beds: one with 50% diluted nutrient solution and one with full concentration nutrient solution.

Figures

Figure 15. Close up of 50% Diluted Nutrient Solution Bed

Figure 16. Close up of Non-Thinned Bed

ResultsThe plants grown in 50% diluted nutrient solution had the best, most vibrant green color

(Figure 15). They had much better color and were healthier looking than the non-thinned plants as well as those grown in full concentration solution. We did not have as many seeds germinate as expected, and many of the seeds in the 50% diluted solution were covered in black fuzz, which we suspected to be mold, also shown in Figure 15. The plants in the full concentration nutrient solution grew decently well, however they were not thriving like those in the diluted solution. They had a yellowish green color and there were also many seeds that were covered in the black fuzz and did not germinate. The plants grown in the non-thinned bed grew well past the points of death of the previous thinned bed. They were yellowed in color which indicates a lack of nutrients (Figure 16). However the fact that they were still alive and growing supports the idea that the thinning was killing the plants and for future plantings we will not thin them. The newly planted seed beds showed that this will work since we only used 1 seed per cap for the beds planted in this experiment.

Problems/potential solutionsThe main problem we encountered was that many of the seeds did not germinate and

were covered in what appeared to be black mold. In order to fix this, we determined that the seedlings should be sterilized before use.

Expt 4 Sterilized seeds

ObjectivesThe first objective of this experiment is to test whether sterilizing the seeds grants us

better success in the germination process. We will sterilize the seeds before stratifying and create new seed beds with the sterilized seeds. The second objective of this experiment is to test a round of plants in the larger system.

Materials and methodsThis experiment followed the same methods as Experiment 3 except using seeds that had

been sterilized before stratifying. The process used for sterilization is shown in Figure 17. We will also only use 1 seed per germination cap and 50% diluted nutrient solution since that was found to be the optimal way to germinate in Experiment 3.

We had enough plants growing at this point that we could move them to the larger system for a round of testing. There were 18 healthy plants that we moved to the larger system shown in Experiment 2. The completed system after the plants had been moved is shown in Figure 19.

Figures

Figure 17. Sterilization Process

Figure 18. Sterilized Seed Bed

Figure 19. Completed Larger System

Results (summarized in tables and described in text)The method of germination with sterilized seeds resulted in 100% of the seeds

germinating (Figure 18). This is a consistent, reliable way to germinate A. thaliana.All but one of the first round of plants tested in the larger system did not survive. We

realized that the larger tubes were sticking up from the solution and that the bottoms of the tubes were touching the bottom of the reservoir causing this. Because of this, the plants were not getting enough water or nutrients since their roots were not long enough.

Problems/potential solutionsTo fix this, we will saw off the bottom of the centrifuge tubes in order to test the newest

batch of seeds.

Expt 5 Larger System with Sawed Centrifuge Tubes

ObjectivesThe objective of this experiment is to test whether sawing off the bottoms of the 50 mL

centrifuge tubes will result in better plant survival and growth. To do this we will saw off the bottoms of each tube at the 10 mL mark and set up the system the same way as Experiment 4.

Materials and methodsThis experiment used the same methods and materials for the larger system in

Experiment 4. The difference is we used a saw to saw off each 50 mL centrifuge tube at the 10 mL mark. Figure 20 shows how the tubes are shorter and are more submerged in the nutrient solution than in the previous experiment. The bottom of each tube is about 2 cm from the bottom of the reservoir. The next two seed beds I planted used seeds that were sterilized before stratifying. The materials and methods were the same as Experiment 4 for germination the seeds. For about two weeks, each seed bed had all the seeds germinate and had healthy looking, green plants. After the second week, considerable algae had begun to grow around the caps and the lid on one of the seed beds. Both beds together had about 40 surviving plants. It was then that we transferred the strongest 35 of those plants to the second iteration of the system designed to grow the larger plants. I used 2 of the 5.5 L reservoirs and had 2 large systems set up to hold all the plants. I have planted beds of sterilized seeds to use for testing in the future while working on optimizing the larger system.

Figures

Figure 20. Completed larger system with sawed off bottoms

Figure 21. Close up of Larger System Surviving Plants

ResultsThe two seed beds began growing very well for the first couple weeks. After the second

week, one of the seed beds began to grow considerable algae and the plants slowly began to die off. The other seed bed was still growing well, however there was slight algal growth the seedlings were slightly yellowed. Once planted in the larger system, many of the plants started to die off in the week after they were planted. There were about 5 that survived and continue to grow, and Figure 21 shows a close up of some of the surviving plants. We realized that many of the roots were not submerged and as such were not getting adequate water or nutrients.

Problems/potential solutionsWe thought that the DI water in the teaching growth chamber might be contributing to

the algae problem. To fix the issue with the algae, we decided it would be best to use sterilized nutrient solution. To do this, we will run the next batch of germination nutrient solution through an autoclave. This should take care of at least part of the algae problem.

For the larger system, we needed a way to address the plants dying. We thought this was mainly due to root desiccation due to the fact that there was a portion of the roots not in the nutrient solution. When inspecting the tubes, we noticed that after transferring, some of the roots wrapped around the threads of the 50 mL centrifuge cap and were not submerged. The way the system is designed leaves a gap where part of the root is not submerged in the water. One potential fix to this issue is to use a thinner sheet of plastic. The layer of plastic isn’t too thick but if it were thinner then there would be less space between the crown of the plant and the nutrient solution. Another potential fix to the problem is to glue the 50 mL centrifuge caps to the bottom of the plastic sheet rather than to the top. That way they would be almost nearly submerged in the nutrient solution.

Final system set-up/conclusionsBased on the results from the previous experiments, one should be able to follow our

procedure to reliably germinate A. thaliana. We were not able to optimize the design of the larger system to have a consistent method of growing larger A. thaliana plants. The issue we ran into is that there is too much space between the crown of each plant and the nutrient solution. The 50 mL centrifuge caps are supposed to be suspended in the solution, but due to the thickness of the cap and the plastic sheet, the entire root is not submerged. This causes the death of many of the plants either because they cannot successfully penetrate the water and stay submerged, or there is too much of the root that is exposed and it undergoes desiccation.

This can be fixed a couple ways. First, we could change the design of larger system we have in place. We could try using a thinner plastic sheet to hold the 50 mL tubes or try gluing the tubes to the bottom of the existing sheet. Both of these changes would let the plant roots sit further in the nutrient solution. Another way we could fix this issue is to change the design of the larger system. To do this, we could create another type of holder that holds the germination caps and keeps the roots submerged in the solution. I think that changing the existing system should be the first step in the next round of testing to determine whether that is a viable option. If it does not work then we need to look into changing the design of the system entirely.

Another thing we noticed was that the 5.5 L reservoirs are relatively shallow. For hydroponics, deeper reservoirs are better because they allow more root growth and their designs allow more options for keeping the roots fully submerged. The first priority should be to find a reliable way to grow larger A. thaliana plants in the larger system. After that, future designs of the larger system should utilize deeper reservoirs to be able to accommodate larger plants. You would also need to change the nutrient solution less often in systems that have deeper reservoirs.

One last thing that we noticed was the odd data regarding the PEG testing in Experiment 1. The data suggests that the PEG used in lab might be contaminated. If this is the case, it will require washings to hopefully remove water soluble contaminants before future use in experiments.

References

Carpita et al. 1979. Determination of the Pore Size of Cell Walls of Living Plant Cells. Science

205: 1144-1147.

Conn et al. 2013. Protocol: optimising hydroponic growth systems for nutritional and

physiological analysis of Arabidopsis thaliana and other plants. Plant Methods 9:4.

He et al. 2014. Endogenous Salicylic Acid Levels and Signaling Positively Regulate Arabidopsis

Response to Polyethylene Glycol-Simulated Drought Stress. Plant Growth Regulation

33:871-880.

Money. 2015. Osmotic Pressure of Aqueous Polyethylene Glycols: Relationship between

Molecular Weight and Vapor Pressure Deficit. Plant Physiology: 1989 766-769.