Embedded Files
Methods
Methods
Safety Protocol:
Gloves, a lab coat, and goggles should always be worn when handling Pseudomonas fluorescens or Bacillus subtilis. If either of these bacteria come into contact with clothing, remove any that is contaminated. If they come into contact with skin, immediately rinse the area with copious amounts of water for several minutes. Refer to the P. fluorescens MSDS and Bacillus subtilis MSDS before handling. When inoculating P. fluorescens and B. subtilis, sterile technique should be used. Sterile technique relies on an open flame to create an updraft and all work services to be sterilized prior to experimentation. Every time a non-flammable chemical is handled along with the bacteria, the mouth and cap of the bottle is flamed before being used and before being closed to keep it sterile. If there are ever any unused or extra cultures of P. fluorescens or B. subtilis, sterilize them using a 5% bleach solution. Do not let the bleach or bleach solution be touched by bare skin, inhaled, drank, or administered to the eyes. If any of this happens immediately rinse the affected area for several minutes or relocate to a place with fresh air. Refer to the Bleach MSDS before handling.
Study Overview:
This study is investigating the effects of Pseudomonas fluorescens (Product # 155256) and Bacillus subtilis (Product # S21890ND) on the growth of red amaranth (Amaranthus cruentus). Use of these rhizobacterial strains was supported by David et al.’s (2018) review on the benefits of Pseudomonas fluorescens as a plant growth promoting rhizobacteria, Ikhajiagbe et al.’s (2021) study on growth responses of amaranthus [L.] species, Cao et al.’s (2023) study on growth responses of rapeseed plants when exposed to microbial consortia, and Devi et al.’s (2022) study on the growth of amaranth (Amaranthus hypochondrius L.). When grown with nitrogen-fixating and mineral solubilizing bacteria strains, all plants showed significantly more growth in the presence of rhizobacteria than no rhizobacteria controls. Although these studies have significantly expanded how beneficial soil bacteria improve plant growth, it is still unknown whether a consortium, or community, of these bacteria could act as a viable and sustainable replacement for chemical fertilizers. The experimental hypothesis is that the amaranth in the presence of a consortium of Pseudomonas fluorescens and Bacillus subtilis, because of the beneficial properties of the two strains of bacteria, will grow significantly larger than the amaranth in the presence of no soil additives and the amaranth receiving the recommended dosage of 20-20-20 NPK chemical fertilizer.
Wisconsin Fast Plant Section:
Wisconsin Fast Plants (Rapid cycling Brassica rapa, Product # B003O3OG82) were used in a screening round to narrow down which combination of Pseudomonas fluorescens (P.f.) and Bacillus subtilis (B.s.) to use with the amaranth. Fast Plants were used in the screening round because of their very high rate of growth and biological similarities to amaranth (Ramos et al., 2021). Fast Plants were divided into 4 groups of 12 pots each, with each group corresponding to different combinations of added bacteria. The first group, the negative control, wasn’t inoculated with any bacteria. The plants from the negative control were expected to grow the least and to act as a baseline for the three experimental arms. Every pot in the second group, the P.f. arm, was inoculated with 1 mL of confluent P. fluorescens. The plants from the P.f. arm were expected to grow more than the no additives arm, the same as the B.s. arm, and worse than the P.f./B.s. arm. Each pot in the third group, the B.s. arm, was inoculated with 1 mL of confluent B. subtilis. The plants from the B.s. arm were expected to grow more than the negative control, the same as the P.f. arm, and worse than the P.f./B.s. arm. Lastly, every pot in the fourth group, the P.f./B.s. arm, was inoculated with 0.5 mL of P. fluorescens and 0.5 mL of B. subtilis. The plants from the P.f./B.s. arm were expected to grow larger than the plants from any other arm. Because of this, it was expected that this combination of bacteria would be used for the experimental arm when growing amaranth.
To begin the trials, two seeds were planted near the center of each pot of every arm and, if both seeds germinated, measurements were averaged. No reference study thinned to less than two plants per pot in their experimentation processes and preliminary data suggests that pots with one plant have no significant advantages compared to pots with two (Cao et al., 2023; Devi et al., 2022; Ikhajiagbe et al., 2021). Because each pot had at least one plant, each arm had no less than twelve biological replicates, meaning that every arm met the requirement of nine biological replicates for statistical analysis.
Fast plants were grown for 30 days, a much shorter time period than Devi et al. (2022) due to the nature of Wisconsin Fast Plants. Since Cao et al. (2023) and Devi et al. (2022) recorded data from their chosen plants throughout their full lifecycle, and the lifecycle of Fast Plants is 30 days, the 30 data collection period was justified for the screening round. Because the bacteria was added to the soil of each pot at confluence, differences in the amount of bacteria in each pot and in each arm were mitigated.
Qualitative plant health observations like plant illness, death, etc. across the 30 days were noted regularly and supplemented with photo checks. Quantitative plant height data the number of leaves on each plant was collected at regular five day intervals. After the 30 day period concluded, final quantitative data of shoot height, number of leaves, and dry mass was gathered and analyzed. Data was analyzed in Google Sheets using two-tailed, independent t-tests. For each set of data, each experimental arm was tested against the negative control and the two arms with the lowest p-values were tested against each other.
Grow Cart / Potting Setup and Planting:
A 7-2-1 mixture of soil was created by mixing 7 lbs of Ocean Forest Potting Soil (Product # B096DT35LS), 2 lbs of Ribbon Organics Compost (Product # B497-000-35BX1), and 1 lb of Biochar (Product # B00Q044LZK) in a plastic container (Product # B07PFLRXN4). The soil was then moistened with 2 L of water. Each seedling pot (Product # B07R9S38VX) in each experimental arm was filled 75% before two Fast Plant seeds were placed 1 cm apart near the center of every pot. The pots were then filled 90%, covering the seeds with a suitable layer of soil for improved root stability and germination rates. Each arm of the experiment was moved to its own seed starter tray (Product # B07R9S38VX) for organization and to avoid cross contamination. Humidity lids (Product # B07R9S38VX) were then placed over each tray to mitigate the possibility that the plants would dry out. Wang et al. (2023) studied germination rates of amaranth seedlings and found this to be an effective method. Although the seedling pots used were much smaller than the ones used by Wang et al. (2023), the same ratio of coverage was used and, since Fast Plants are much smaller than amaranth, the difference was negligible.
The top shelf of the grow cart (Product # BA3P2) was chosen as the place to grow the plants for a multitude of reasons. First, the spacing between the trays and lights are adjustable, which allowed the top section of the cart to have 20 inches of vertical room for the plants to grow. On average, Fast Plants grow to around 6 in, so this was more than enough space. Secondly, the plug that corresponded to the light at the top of the cart was easily accessible and, therefore, it was easy to set up the programmable light timer (Product # B094PFJ5JW). The timer was set to turn on the light eight hours per day at high intensity so that the plants all got an even and suitable amount of light to conduct photosynthesis. This setup was also used by Sadvakasova et al. (2023) in their study about a synthetic algocyanobacterial consortium and by Tyagi et al. (2023) in their study testing whether rhizobacteria could boost drought resistance in maize.
Labeling tape (Product # B0889LYH61) and a black sharpie (Product # B00G4CJ8GK) were used to label each arm of the experiment as well as each seedling pot. The humidity lids for each arm’s respective seed starter tray were taken off and placed in a safe area. The seed starter tray for the negative control was labeled ‘Neg’ and each of the seedling pots were labeled ‘Neg 1’ - ‘Neg 12’ respectively. The seed starter tray for the P.f. experimental arm was labeled ‘P.f.’ and each of the seedling pots were labeled ‘P.f. 1’ - ‘P.f. 12’ respectively. The seed starter tray for the B.s. experimental arm was labeled ‘B.s.’ and each of the seedling pots were labeled ‘B.s. 1’ - ‘B.s. 12’ respectively. The seed starter tray for the P.f./B.s. experimental arm was labeled ‘P.f./B.s.’ and each of the seedling pots were labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’ respectively. The humidity lids for each arm were then placed back on their respective seed starter trays.
Bacterial Culturing and Soil Inoculation:
Refer to the Safety Protocol section before continuing and make sure to wear gloves, a lab coat, and safety goggles. Two 15 mL transformation tubes (Product # 215090) were labeled ‘P.f.’ using labeling tape and a black sharpie. Using bleach (Product # B01C65T4M6), tap water, and a 250 mL beaker (Product # B08D6W1Z4S), an approximately 5% bleach solution was made in case there was any excess P. fluorescens culture that had to be sterilized. The bunsen burner (Product # B085QLY8C8) was plugged into the gas hose and maneuvered so that it wasn’t under anything lower than 4 feet above it. This was to make sure it wouldn’t unintentionally set anything on fire. The gas was then turned on, the lighter (Product # B08BQXM2RT) was used to light the gas, and it was made sure that the flame was blue and created a triangle near the top. The bunsen burner acted as a precaution to uphold the sterile technique mentioned in the Safety Protocol. All materials were moved within a 6 in radius of the lit bunsen burner. Within that radius, the serological pipette (Product # B09KGXTVBF) with a sterile 5 mL end (Product # B0C1S2N4NZ) was used to pipette 12 mL of nutrient broth (Product # 776380) into each labeled transformation tube. Refer to the Nutrient Broth MSDS before handling. The Pseudomonas fluorescens plate culture (Product # 155256) was opened and a sterile inoculation loop (Product # B079TNFHT4) was used to transfer a swabbing of Pseudomonas fluorescens into each transformation tube. The amounts swabbed were approximate because all measurements involving inoculating the soil with the bacteria were made while the culture was at confluence. This method is supported by Devi et al. (2022), Ikhajiagbe et al. (2021), and Negi et al. (2022). The lids of the tubes were closed and the plate of Pseudomonas fluorescens was stored in the fridge. The tubes labeled ‘P.f.’ were then placed into the incubator while other tubes were prepared. This process was then repeated to make two Bacillus subtilis cultures in two tubes labeled ‘B.s.’ using the slant culture of Bacillus subtilis (Product # S21890ND). All materials except for the tubes labeled ‘P.f.’ and ‘B.s.’ were cleaned up or thrown away. The tubes labeled ‘P.f.’ and ‘B.s.’ were then placed inside the shaking incubator (Product # 10000070759) so that they could grow to confluence (Negi et al., 2022). The shaking incubator was subsequently set to run at 30 ℃, at 250 rpm, for 24 hours (Devi et al., 2022).
The next day, safety equipment was put back on. The cultures had reached confluence and were taken out of the shaking incubator so that they could be used to make glycerol stocks to facilitate future experimentation. Ten 2 mL screw cap vials (Product # B0CGT6T2S6) were labeled ‘P.f. 1’ - ‘P.f. 10’ and ten 2 mL screw cap vials were labeled ‘B.s. 1’ - ‘B.s. 10’. Using tap water, glycerol (Product # B00DYODWHC), and a graduated cylinder, a 50% water 50% glycerol solution was made. Refer to the glycerol MSDS before handling. The bleach solution was brought over in case there was any excess P. fluorescens or Bacillus subtilis culture that had to be sterilized. The bunsen burner was plugged into the gas hose and maneuvered so that it wasn’t under anything lower than 4 feet above it. This was to make sure it wouldn’t unintentionally set anything on fire. The gas hose was then turned on, the lighter was used to light the gas, and it was made sure that the flame was blue and created a triangle near the top. The bunsen burner acted as a precaution to uphold the sterile technique mentioned in the Safety Protocol. All materials were moved within a 6 in radius of the lit bunsen burner. Still within that radius, the 5 mL serological pipette was used to pipette 1 mL of the 50-50 glycerol-water solution into every 2 mL screw cap tube. The end of the serological pipette was discarded and a new, sterile end was inserted. The P1000 micropipette was then used to pipette 1 mL of Pseudomonas fluorescens culture from one of the transformation tubes labeled ‘P.f.’ into each of the 2 mL screw cap vials labeled ‘P.f. 1’ - ‘P.f. 10’. The end of the serological pipette was discarded and a new, sterile end was inserted. The P1000 micropipette was then used to pipette 1 mL of Bacillus subtilis culture from one of the transformation tubes labeled ‘B.s.’ into each of the 2 mL screw cap vials labeled ‘B.s. 1’ - ‘B.s. 10’. The Safety Protocol was reviewed and the excess cultures of Bacillus subtilis and Pseudomonas fluorescens from the used tubes were sterilized using the 5% bleach solution. All of the 2 mL screw caps were then stored in the freezer for future use. All materials except for the bleach solution and the two remaining tubes, one labeled ‘P.f.’ and one labeled ‘B.s.’, were cleaned up or thrown away to clear space.
The two remaining tubes, one labeled ‘P.f.’ and one labeled ‘B.s.’, were then used to inoculate the soil of the experimental arms. The seed starter trays labeled ‘P.f.’, ‘B.s.’, & ‘P.f./B.s.’ were brought to a large workspace. The humidity lids for each arm’s respective seed starter tray were taken off and placed in a safe area. The P1000 micropipette (Product # BT1503) with a sterile end was used to evenly distribute 0.5 mL of Pseudomonas fluorescens culture into each of the seedling pots labeled ‘P.f. 1’ - ‘P.f. 12’. The same pipette ending was then used to evenly distribute 0.25 mL of Pseudomonas fluorescens culture into each of the seedling pots labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’. This method of inoculation is supported by De Salamone et al.’s (2012) study on the impact of plant genotypes on rhizosphere microbial communities and field crop production. The end of the serological pipette was then discarded and replaced with a new, sterile one. The 5 mL serological pipette with a sterile end was used to evenly distribute 0.5 mL of Bacillus subtilis culture into each of the seedling pots labeled ‘B.s. 1’ - ‘B.s. 12’. The same pipette ending was then used to evenly distribute 0.25 mL of Bacillus subtilis culture into each of the seedling pots labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’ (De Salamone et al., 2012). The humidity lids for each arm were then placed back on their respective seed starter trays. All seed starter trays were then moved back into their designated area of the top shelf of the grow cart. The 5% bleach solution was subsequently used to sterilize the remaining Pseudomonas fluorescens and Bacillus subtilis cultures. All materials were cleaned up or thrown away.
Plant Care and Watering:
Every plant in each arm was watered with 150 mL of tap water every 3 days. This was measured with a graduated cylinder. This method of watering is supported by Aqeel et al.'s (2023) review about how plant-soil-microbe interactions maintain ecosystem stability and coordinate turnover under constantly changing environmental conditions. The plants and, in turn, the seed starter trays for each arm of the experiment were kept inside of the grow cart and under their respective humidity lids at all times. This was to make sure that the plants remained within the temperature confines of 21 ℃ - 27 ℃ at all times and to make sure that the soil didn’t lose too much water in between watering sessions.
Data Collection and Analysis:
After a 15 day growing period had passed, all seed starter trays were removed from the grow cart and placed onto a large, open workspace. The humidity lids for each arm’s respective seed starter tray were taken off and placed in a safe area. A ruler was then used to measure the height, in centimeters, of the Fast Plant growing in the seedling container labeled ‘Neg 1’. Height was quantified as the tallest point of the plant above the soil. The number of leaves of the plant in the seedling pot labeled ‘Neg 1’ was counted. The height and number of leaves were subsequently recorded in their corresponding cells of a Google Sheets page that was used to analyze quantitative data. This process was then repeated for the plants in the seedling pots labeled ‘Neg 2’ - ‘Neg 12’ and all plants in all experimental arms. If there was more than one plant in a seedling pot, both plants were measured and the average value of the measurements was inputted into Google Sheets. If there was no plant in a given pot, all data regarding that pot was omitted following the supposed death of the plant. This entire process was repeated at the 20, 25, and 30 day growing points.
Once all of the data about height and number of leaves was entered into the Google Sheets page from the 15, 20, 25, and 30 day checkpoints, it was centralized to facilitate easier data analysis. The values of height for every plant in the negative control at the 15 day mark were averaged in a new cell. This process was completed for the 20, 25, and 30 day marks as well. This was repeated for the ‘P.f.’ arm, the ‘B.s.’ arm, and the ‘P.f./B.s.’ arm. This entire process was repeated for data about the number of leaves on each plant. The averages of the height data of the ‘P.f.’ arm, the ‘B.s.’ arm, and the ‘P.f./B.s.’ arm at the 30 day checkpoint were individually compared to the average height data of the negative control after 30 days using a two-tailed, independent t-test to measure the significance of their respective differences. The two experimental arms with the lowest p-value (the cutoff being a p-value of less than 0.05), and therefore the highest significant difference from the negative control, were compared using a two-tailed, independent t-test to measure the significance of their difference. The same process was subsequently completed for data representing the number of leaves on a given plant. The hypothesis would be supported if the plants in the P.f./B.s. arm grew significantly more leaves than plants in the other arms.
Using a black sharpie, 49 weighing boats (Product # …) were labeled to facilitate dry mass data collection. The insides of 12 of the weighing boats were labeled ‘Neg 1’ - ‘Neg 12’, 12 were labeled ‘P.f. 1’ - ‘P.f. 12’, 12 were labeled ‘B.s. 1’ - ‘B.s. 12’, and 12 were labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’. The plants were then delicately uprooted and placed in the weighing boat that corresponded to their arm and number. All weighing boats with plants were placed into the dehydrator (Product # B09J97F7L9), which was turned to its medium setting, for 12 hours. This process took place overnight. The weighing boats were then taken out of the dehydrator and placed in an open workspace near the analytical balance (Product # B08YN7HWSQ). The balance was turned on and the extra, empty weighing boat was placed onto the scale. The balance was then zeroed so that it didn’t account for the weight of the weighing boat when measuring the dry mass of the plants. The weighing boat with the dehydrated plant labeled ‘Neg 1’ was then placed on the scale of the balance and the weight displayed was recorded in the corresponding section of the Google Sheets page in grams. This process was then repeated for all other labeled weighing boats (‘Neg 2’ - ‘Neg 12’, ‘P.f. 1’ - ‘P.f. 12’, ‘B.s. 1’ - ‘B.s. 12’, and ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’). The dry masses of each arm were then respectively averaged in separate cells of the Google Sheet. The averages of the dry mass data of the ‘P.f.’ arm, the ‘B.s.’ arm, and the ‘P.f./B.s.’ arm were individually compared to the average dry mass data of the negative control using a two-tailed, independent t-test to measure the significance of their respective differences. Subsequently, the two experimental arms with the lowest p-value, and therefore the highest significant difference from the negative control, were compared to each other using a two-tailed, independent t-test to measure the significance of their difference. The outcome that would support the hypothesis would be that the plants grown in the P.f./B.s. arm would be significantly heavier than plants from other arms.
Amaranth Section:
Amaranth (Amaranthus cruentus, Product # B005ES9LU8) was used to test if a consortium of Pseudomonas fluorescens (P.f.) and Bacillus subtilis (B.s.) could grow amaranth better than 20-20-20 NPK chemical fertilizer (Product # B001REA5NK). Amaranth was used because of its potential as a substitute for grains that require significantly more water. Amaranth was divided into 3 groups of 9 pots each, with each group corresponding to different soil additives. The first group, the negative control, wasn’t inoculated with any bacteria. The plants from the negative control were expected to grow the least and to act as a baseline for the two other arms. Every pot in the second group, the fertilizer arm, was watered with the 100% recommended dosage of 20-20-20 NPK chemical fertilizer. The plants from the fertilizer arm were expected to grow more than the no additives arm and worse than the P.f./B.s. arm. Lastly, every pot in the third group, the P.f./B.s. arm, was inoculated with 0.5 mL of P. fluorescens and 0.5 mL of B. subtilis. The plants from the P.f./B.s. arm were expected to grow larger than the plants from the other arms. Because of this, it was expected that this combination of bacteria would be used for the experimental arm when growing amaranth.
To begin the trials, two seeds were planted near the center of each pot of every arm and, if both seeds germinated, measurements were averaged. No reference study thinned to less than two plants per pot in their experimentation processes and preliminary data suggests that pots with one plant have no significant advantages compared to pots with two (Cao et al., 2023; Devi et al., 2022; Ikhajiagbe et al., 2021). Because each pot had at least one plant, each arm had no less than nine biological replicates, meaning that every arm met the requirement of nine biological replicates for statistical analysis.
Amaranth was grown for 40 days, a much shorter time period than Devi et al. (2022) due to time constraints. Since Cao et al. (2023) and Devi et al. (2022) recorded data from their chosen plants at the 45 day mark, and the plants were measured after 40 days, the 40 day data collection period was justified for the experiment. Because the bacteria was added to the soil of each pot at confluence, differences in the amount of bacteria in each pot were mitigated.
Qualitative plant health observations like plant illness, death, etc. across the 40 days were noted regularly and supplemented with photo checks. Quantitative plant height data the number of leaves on each plant was collected at regular five day intervals. After the 40 day period concluded, final quantitative data of shoot height and number of leaves was gathered and analyzed. Data was analyzed in Google Sheets using two-tailed, independent t-tests. For each set of data, each arm was tested against each other arm.
Grow Cart / Potting Setup and Planting:
A 7-2-1 mixture of soil was created by mixing 7 lbs of Ocean Forest Potting Soil (Product # B096DT35LS), 2 lbs of Ribbon Organics Compost (Product # B497-000-35BX1), and 1 lb of Biochar (Product # B00Q044LZK) in a plastic container (Product # B07PFLRXN4). The soil was then moistened with 2 L of water. Each seedling pot (Product # B07R9S38VX) in each experimental arm was filled 75% before two amaranth seeds were placed 1 cm apart near the center of every pot. The pots were then filled 90%, covering the seeds with a suitable layer of soil for improved root stability and germination rates. Each arm of the experiment was moved to its own seed starter tray (Product # B07R9S38VX) for organization and to avoid cross contamination. Humidity lids (Product # B07R9S38VX) were then placed over each tray to mitigate the possibility that the plants would dry out. Wang et al. (2023) studied germination rates of amaranth seedlings and found this to be an effective method. The seedling pots used were slightly smaller than the ones used by Wang et al. (2023), but the same ratio of coverage was used.
The top shelf of the grow cart (Product # BA3P2) was chosen as the place to grow the plants for a multitude of reasons. First, the spacing between the trays and lights was adjustable, which allowed the top section of the cart to have 20 inches of vertical room for the plants to grow. On average, amaranth can only get to around 10 in, so this was more than enough space. Secondly, the plug that corresponded to the light at the top of the cart was easily accessible and, therefore, it was easy to set up the programmable light timer (Product # B094PFJ5JW). The timer was set to turn on the light eight hours per day at high intensity so that the plants all got an even and suitable amount of light to conduct photosynthesis. This setup was also used by Sadvakasova et al. (2023) in their study about a synthetic algocyanobacterial consortium and by Tyagi et al. (2023) in their study testing whether rhizobacteria could boost drought resistance in maize.
Labeling tape (Product # B0889LYH61) and a black sharpie (Product # B00G4CJ8GK) were used to label each arm of the experiment as well as each seedling pot. The humidity lids for each arm’s respective seed starter tray were taken off and placed in a safe area. The seed starter tray for the negative control was labeled ‘Neg’ and each of the seedling pots were labeled ‘Neg 1’ - ‘Neg 9’ respectively. The seed starter tray for the fertilizer experimental arm was labeled ‘Fert’ and each of the seedling pots were labeled ‘Fert 1’ - ‘Fert 9’ respectively. The seed starter tray for the P.f./B.s. experimental arm was labeled ‘P.f./B.s.’ and each of the seedling pots were labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 9’ respectively. The humidity lids for each arm were then placed back on their respective seed starter trays.
Bacterial Culturing and Soil Inoculation:
Refer to the Safety Protocol section before continuing and make sure to wear gloves, a lab coat, and safety goggles. Two 15 mL transformation tubes (Product # 215090) were labeled ‘P.f.’ using labeling tape and a black sharpie. Using bleach (Product # B01C65T4M6), tap water, and a 250 mL beaker (Product # B08D6W1Z4S), an approximately 5% bleach solution was made in case there was any excess P. fluorescens culture that had to be sterilized. The bunsen burner (Product # B085QLY8C8) was plugged into the gas hose and maneuvered so that it wasn’t under anything lower than 4 feet above it. This was to make sure it wouldn’t unintentionally set anything on fire. The gas was then turned on, the lighter (Product # B08BQXM2RT) was used to light the gas, and it was made sure that the flame was blue and created a triangle near the top. The bunsen burner acted as a precaution to uphold the sterile technique mentioned in the Safety Protocol. All materials were moved within a 6 in radius of the lit bunsen burner. Within that radius, the serological pipette (Product # B09KGXTVBF) with a sterile 5 mL end (Product # B0C1S2N4NZ) was used to pipette 12 mL of nutrient broth (Product # 776380) into each labeled transformation tube. Refer to the Nutrient Broth MSDS before handling. The Pseudomonas fluorescens plate culture (Product # 155256) was opened and a sterile inoculation loop (Product # B079TNFHT4) was used to transfer a swabbing of Pseudomonas fluorescens into each transformation tube. The amounts swabbed were approximate because all measurements involving inoculating the soil with the bacteria were made while the culture was at confluence. This method is supported by Devi et al. (2022), Ikhajiagbe et al. (2021), and Negi et al. (2022). The lids of the tubes were closed and the plate of Pseudomonas fluorescens was stored in the fridge. The tubes labeled ‘P.f.’ were then placed into the incubator while other tubes were prepared. This process was then repeated to make two Bacillus subtilis cultures in two tubes labeled ‘B.s.’ using the slant culture of Bacillus subtilis (Product # S21890ND). All materials except for the tubes labeled ‘P.f.’ and ‘B.s.’ were cleaned up or thrown away. The tubes labeled ‘P.f.’ and ‘B.s.’ were then placed inside the shaking incubator (Product # 10000070759) so that they could grow to confluence (Negi et al., 2022). The shaking incubator was subsequently set to run at 30 ℃, at 250 rpm, for 24 hours (Devi et al., 2022).
The next day, safety equipment was put back on. The cultures had reached confluence and were taken out of the shaking incubator so that they could be used to make glycerol stocks to facilitate future experimentation. Ten 2 mL screw cap vials (Product # B0CGT6T2S6) were labeled ‘P.f. 1’ - ‘P.f. 10’ and ten 2 mL screw cap vials were labeled ‘B.s. 1’ - ‘B.s. 10’. Using tap water, glycerol (Product # B00DYODWHC), and a graduated cylinder, a 50% water 50% glycerol solution was made. Refer to the glycerol MSDS before handling. The bleach solution was brought over in case there was any excess P. fluorescens or Bacillus subtilis culture that had to be sterilized. The bunsen burner was plugged into the gas hose and maneuvered so that it wasn’t under anything lower than 4 feet above it. This was to make sure it wouldn’t unintentionally set anything on fire. The gas hose was then turned on, the lighter was used to light the gas, and it was made sure that the flame was blue and created a triangle near the top. The bunsen burner acted as a precaution to uphold the sterile technique mentioned in the Safety Protocol. All materials were moved within a 6 in radius of the lit bunsen burner. Still within that radius, the P1000 micropipette was used to pipette 1 mL of the 50-50 glycerol-water solution into every 2 mL screw cap tube. The end of the serological pipette was discarded and a new, sterile end was inserted. The P1000 micropipette was then used to pipette 1 mL of Pseudomonas fluorescens culture from one of the transformation tubes labeled ‘P.f.’ into each of the 2 mL screw cap vials labeled ‘P.f. 1’ - ‘P.f. 10’. The end of the serological pipette was discarded and a new, sterile end was inserted. The P1000 micropipette was then used to pipette 1 mL of Bacillus subtilis culture from one of the transformation tubes labeled ‘B.s.’ into each of the 2 mL screw cap vials labeled ‘B.s. 1’ - ‘B.s. 10’. The Safety Protocol was reviewed and the excess cultures of Bacillus subtilis and Pseudomonas fluorescens from the used tubes were sterilized using the 5% bleach solution. All of the 2 mL screw caps were then stored in the freezer for future use. All materials except for the bleach solution and the two remaining tubes, one labeled ‘P.f.’ and one labeled ‘B.s.’, were cleaned up or thrown away to clear space.
The two remaining tubes, one labeled ‘P.f.’ and one labeled ‘B.s.’, were then used to inoculate the soil of the experimental arms. The seed starter tray labeled ‘P.f./B.s.’ was brought to a large workspace. The humidity lid for the arm’s seed starter tray was taken off and placed in a safe area. The P1000 micropipette (Product # BT1503) with a sterile end was used to evenly distribute 0.5 mL of Pseudomonas fluorescens culture into each of the seedling pots labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’. This method of inoculation is supported by De Salamone et al.’s (2012) study on the impact of plant genotypes on rhizosphere microbial communities and field crop production. The end of the serological pipette was then discarded and replaced with a new, sterile one. The 5 mL serological pipette with a sterile end was used to evenly distribute 0.5 mL of Bacillus subtilis culture into each of the seedling pots labeled ‘P.f./B.s. 1’ - ‘P.f./B.s. 12’ (De Salamone et al., 2012). The humidity lid for the arm was then placed back on its seed starter tray. The seed starter tray was then moved back into its designated area of the top shelf of the grow cart. The 5% bleach solution was subsequently used to sterilize the remaining Pseudomonas fluorescens and Bacillus subtilis cultures. All materials were cleaned up or thrown away.
Plant Care and Watering:
Every plant in each arm was watered with 150 mL of tap water every 3 days. This was measured with a graduated cylinder. This method of watering is supported by Aqeel et al.'s (2023) review about how plant-soil-microbe interactions maintain ecosystem stability and coordinate turnover under constantly changing environmental conditions. The plants and, in turn, the seed starter trays for each arm of the experiment were kept inside of the grow cart and under their respective humidity lids at all times. This was to make sure that the plants remained within the temperature confines of 21 ℃ - 27 ℃ at all times and to make sure that the soil didn’t lose too much water in between watering sessions.
Data Collection and Analysis:
After a 10 day growing period had passed, all seed starter trays were removed from the grow cart and placed onto a large, open workspace. The humidity lids for each arm’s respective seed starter tray were taken off and placed in a safe area. A ruler was then used to measure the height, in centimeters, of the Fast Plant growing in the seedling container labeled ‘Neg 1’. Height was quantified as the tallest point of the plant above the soil. The number of leaves of the plant in the seedling pot labeled ‘Neg 1’ was counted. The height and number of leaves were subsequently recorded in their corresponding cells of a Google Sheets page that was used to analyze quantitative data. This process was then repeated for the plants in the seedling pots labeled ‘Neg 2’ - ‘Neg 12’ and all plants in all experimental arms. If there was more than one plant in a seedling pot, both plants were measured and the average value of the measurements was inputted into Google Sheets. If there was no plant in a given pot, all data regarding that pot was omitted following the supposed death of the plant. This entire process was repeated at the 20, 30, and 40 day growing points.
Once all of the data about height and number of leaves was entered into the Google Sheets page from the 10, 20, 30, and 40 day checkpoints, it was centralized to facilitate easier data analysis. The values of height for every plant in the negative control at the 10 day mark were averaged in a new cell. This process was completed for the 20, 30, and 40 day marks as well. This was repeated for the ‘Fert’ arm and the ‘P.f./B.s.’ arm. This entire process was repeated for data about the number of leaves on each plant. The averages of the height data of the ‘Fert’ arm and the ‘P.f./B.s.’ arm at the 40 day checkpoint were individually compared to the average height data of the no additives arm after 40 days using a two-tailed, independent t-test to measure the significance of their respective differences. The two experimental arms with the lowest p-value (the cutoff being a p-value of less than 0.05), and therefore the highest significant difference from the negative control, were compared using a two-tailed, independent t-test to measure the significance of their difference. The same process was subsequently completed for data representing the number of leaves on a given plant. The hypothesis would be supported if the plants in the P.f./B.s. arm grew significantly more leaves than plants in the other arms.
Materials List and Procedure (Click to View):
Materials List and Procedure (Click to View):
Materials List:
Pseudomonas fluorescens Living Plate & Pseudomonas fluorescens MSDS
Petri Dishes, Polystyrene, Disposable, Sterile, 60 x 15 mm, Pack of 20
Containers (Kingrol 24 pack of Mini Clear Plastic Containers) 7.3 x 7.3 x 2.5 Centimeter
Refrigerator
Water
Seed Starter with lid
Computer to record data
Macbook or chromebook is preferred
Phone (Android/iPhone preferred)
Greenhouse Setup:
Find an area for the grow cart that is spacious, close to an outlet, and is within the general temperature range of 21 ℃ - 28 ℃. Proceed to roll the grow cart to this area.
Clear the trays of the grow cart of any extra materials and put them in the plastic organizer bins. Store the bins in an empty section of a cabinet.
Using three sections of 7 cm masking tape and the black sharpie, label the three sub-trays in the bottom racktray ‘Neg Section’, ‘Pos Section’, and ‘Exp Section’ respectively.
Using the ruler, from the base of the bottom tray, measure 76 cm up the side of the cart and make a mark using the sharpie.
If there are any trays in between the floor of the first racktray and the sharpie mark, use the adjustable wrench to unscrew the bolts of the shelf(s) and push it/them to the bottom. Make sure to hold onto the shelf as it’s being unscrewed because it may fall without support.
Using the knob on the side of the light fixture, move it up until the bottom of it is approximately 1 cm above the mark.
Place one of the digital thermometers into the middle tray so that it's readable and its stand is holding it up.
Clear a workspace next to the grow cart where it can be plugged in.
Plug in the 2nd lowest light on the grow cart into the back of one of the programmable timers and, using an extension cord, plug in the side of the timer into an outlet.
Once the digital display is active for 5 minutes, test that the light is turning on by pressing the ON/AUTO/OFF button until the bottom right of the display reads ‘on’.
If the light isn’t working, try plugging the timer into a different outlet or switching the extension cord out for the other one.
The illumination of the red light above the display signifies that the timer is getting power so, if the light isn’t working then, try checking the connection.
Once the light is on, press the ON/AUTO/OFF button until the display reads ‘off’ to turn off the light and then press it once more so the light is off and the display reads ‘auto’.
Use the phone as a reference for what time of the day and day of the week it is for the rest of this procedure.
Hold the CLOCK button until the end of step 15.
Press the WEEK button until the top of the display reads what day of the week it is.
Press the HOUR button until the center of the display reads what hour it is and the top left of the display reads either ‘AM’ or ‘PM’ depending on the time of day.
Press the MIN button until the time displayed on the timer perfectly matches up with the time displayed on the phone.
Press the PROG button once.
This is to set the starting time for the light to turn on.
Press the WEEK button once so that all days of the week are displayed on the top of the screen.
Press the HOUR button until the timer reads ‘8:00 am’.
Press the MIN button until the timer reads ‘8:30 am’.
Press the PROG button once.
This is to set the time for the light to turn off.
Press the WEEK button once so that all days of the week are displayed on the top of the screen.
Press the HOUR button until the timer reads ‘4:00 pm’.
Press the MIN button until the timer reads ‘4:30 pm’.
Press the CLOCK button once to get back to the normal view.
If anything went wrong, use the pencil to press the small circular button on the right of the timer to reset the timer. After this, wait 5 minutes and repeat steps 7-19.
Clean up all materials and make sure that the display of the timer isn’t on the ground to avoid any accidental changes.
Fast Plant Screening:
General Care Info for Fast Plants & Microbial Consortiums:
Plant Section:
Seed Section:
Using a ruler, measure a 5 cm section of masking tape and use it to label a 7.3 cm x 7.3 cm container “Fast Plant Seed Storage” with a black sharpie. Make sure to not cover the lid so the container can open.
The fast plant seeds should be stored in the fridge, as they keep best between 2 ℃ and 4 ℃.
Will keep at least a year if stored this way.
If the seeds are permitted to be at room temperature and exposed to moisture, they have the possibility of sprouting, ruining the experiment.
Ready the Fast Plant Seed Storage’ container (will be referred to as seed container) by wiping it down with a section of paper towel so that it is completely dry, and place all of the seeds into it.
Place the seed container in the fridge when the seeds aren’t being used to ensure that germination doesn’t occur.
Take the “Fast Plant Seed Storage Container” out of the fridge and place it on the workstation.
Cut a piece of the paper towel so that, when folded into fourths, it fits in the 4 oz jars
Open the fast plant seed container and take eighteen seeds out.
Fill the 250 mL beaker with tap water so that it’s halfway filled and dip a section of the cut paper towel in so it’s completely wet.
This wetted paper towel section will be used to pre-germinate the seeds to ensure only viable seeds are planted in the soil at the beginning of the experiment.
Set out the paper towel section flat on the workspace, place all of the seeds around the centerline of the width of the paper towel, each approximately ½ cm away from each other. Fold the towel into fourths with the first fold being vertically and the second being horizontally.
Place the folded and wetted paper towel section into one of the 4 oz jars and close the jar.
This is so the paper towel stays wet and the seeds can germinate.
Using a piece of appropriately sized masking tape and a sharpie, label the jar “Neg”
Place the jar into the incubator and set the temperature to 23 ℃. If this is consistently around room temperature, just leave the jars out of the incubator.
Repeat steps 9-15 three more times to create another seven sets of eighteen seeds for germination except, when doing step 7, label the jar “P. f.” the second time, “B. s.” the third time, and “P. f. & B. s.” the fourth time.
Again, this is to ensure that there are at least nine viable germinated seeds for the experiment.
Using the stand on its back to stand it up, place one of the digital thermometers into the incubator.
This will help as it displays the temperature that the seeds are actually at.
Close and place the seed storage container back into the fridge.
Any seeds that don’t germinate after 18 days of being in the jar should be placed into the garbage (there may still be extra germinated seeds in the jar to ensure that all sections of the procedure have enough germinated seeds for planting). If there are no more seeds in the jar, throw away the paper towel and place the jar back into storage.
Temperature & Watering Section:
After the seeds have germinated and are planted (described in Seed Subsection steps 4-14 & Negative Control steps 11-20), using a 1 L graduated cylinder to measure, water each plant with 500 mL of tap water every three days.
Keep plants / the grow cart inside at all times as the temperature should always be between 21 ℃ and 27 ℃. If the temperature is ever outside the range, investigate why this may be the case and, if necessary, move the grow cart to a cooler location or open the flaps of the humidity tent.
Any temperature outside of these parameters could affect the growth of the plants and, therefore, the experiment.
Note that amaranth does not have any specific humidity requirements.
To ensure that the temperature is always in the range, every time the plants are watered, check the digital thermometer.
P.f./B.s. Culturing Section:
Put on nitrile gloves, goggles, and a lab coat.
Using a ruler to measure, cut twenty sections of 5 cm by 2.5 cm masking tape.
Halfway down an unlabeled 15 mL transformation tube, stick on one of the previously made sections of tape horizontally to make a label. Do this for the nineteen remaining tape sections and unlabelled 15 mL transformation tubes.
Using the black sharpie, label the tape on the twenty tubes “P.f. 1” - “P.f. 20” respectively so each tube is assigned an experimental arm designation and unique number.
Using the bleach, tap water, and the 250 mL beaker, make a 5% bleach solution by adding 10 mL of bleach and 190 mL of tap water to the beaker. Do not let the bleach or bleach solution be touched by bare skin, inhaled, drank, or administered to the eyes. If any of this happens immediately rinse the affected area for several minutes or relocate to a place with fresh air.
Refer to the bleach MSDS before handling.
Plug the bunsen burner into the gas hose, turn on the gas, and use a lighter to light the flame. Make sure the burner isn’t immediately under anything and that the flame is blue and makes a triangle at its peak.
This is so all materials remain sterile as long as they’re close to the flame (due to the hot air rising and taking foreign bacteria with it).
Additionally, any non-flammable material (such as caps, lips of bottles, and inoculation loops) can be sterilized by running them through the flame.
Move all materials within a 6 inch radius of the burner.
Using the 50 mL serological pipette, fill each tube with the nutrient broth up to the 9 mL mark. Keep the broth away from any open flames, any consumables, and skin. If it touches skin, immediately rinse the area thoroughly.
Refer to the nutrient broth MSDS before handling.
Place the Pseudomonas fluorescens (P. fluorescens) nutrient agar petri dish culture on a flat surface and open it using the scissors. Don’t let the P. fluorescens get onto any wearables or skin. If it comes into contact with clothing, remove any that is contaminated. If it comes into contact with skin, immediately rinse the area with copious amounts of water for several minutes.
Refer to the P. fluorescens MSDS before handling.
Using a sterile inoculation loop, transfer a small swabbing of the original P. fluorescens from the petri dish into the tube labeled ‘P.f. 1’. Repeat this process for tubes ‘P.f. 2’ - ‘P.f. 20’.
The value of scraped P. fluorescens is approximate because, after it is made into a liquid culture and is grown, there will be precise measurements for how much to put into each plant from the Experimental Control section.
Additionally, there isn’t a large amount of bacteria because too much initial P. fluorescens can lead to the death of the culture.
Any excess P. fluorescens from the petri dish culture should be stored in the fridge at 4 ℃. Once this is done, store the agar in the fridge, wash the petri dish out using the 5% bleach solution, and place it back with other materials. In addition, wash the scoopula out using the 5% bleach solution and place it back with other materials.
Using the cap, close the tubes so the solution doesn’t fall out.
Take the tubes over to the shaking incubator, make sure it’s on, open it, and place the tubes in the holder. Make sure to close the shaking incubator.
Set the temperature of the shaking incubator to 35 ℃ by holding down the central knob until the light next to the ‘temperature’ setting starts blinking and then turning it until the display displays the desired value.
and the rpm of the shaking incubator to 250 rpm by holding down the central knob until the light next to the ‘rpm’ setting starts blinking and then turning it until the display displays the desired value.
Store any miscellaneous materials (i.e. masking tape, ruler, sharpie, etc.) and PPE back with other materials.
This is a good place to pause as the next step involves a large amount of waiting time.
Set the shaking incubator run for 18 hours by holding down the central knob until the light next to the ‘time’ setting starts blinking and then turning it until the display displays the desired value so the P. fluorescens can mix completely with the nutrient broth. Press the ‘start’ button.
The run time should take place overnight.
Put all PPE back on.
Once the 18 hours have passed, remove all tubes from the shaking incubator, place them in the fridge (4 ℃). The cultures can survive for a month in these conditions, but they will only have to be in there long enough for the seeds to germinate (less than 18 days).
At the end of this process all labeled tubes should contain a liquid medium of nutrient broth and P. fluorescens and be in the fridge.
Repeat this process twice, except replace any instances of P. fluorescens with B. subtilis the first time and any instances of P. fluorescens with A. chroococcum the second time.
MSDS for all bacteria is in the materials list.
General Care Info for Amaranth & Microbial Consortium (References Amaranth & MC):
Plant Section:
Seed Section:
Using a ruler, measure a 5 cm section of masking tape and use it to label a 7.3 cm x 7.3 cm container “Amaranth Seed Storage” with a black sharpie. Make sure to not cover the lid so the container can open.
The amaranth seeds should be stored in the fridge, as they keep best between 2 ℃ and 4 ℃.
Will keep at least a year if stored this way.
If the seeds are permitted to be at room temperature and exposed to moisture, they have the possibility of sprouting, ruining the experiment.
Ready the ‘Amaranth Seed Storage’ container (will be referred to as seed container) by wiping it down with a section of paper towel so that it is completely dry, and place all of the seeds into it.
Place the amaranth seed container in the fridge when the seeds aren’t being used to ensure that germination doesn’t occur.
Take the “Amaranth Seed Storage Container” out of the fridge and place it on the workstation.
Cut a piece of the paper towel so that, when folded into fourths, it fits in the 4 oz jars
Open the seed container and take eighteen seeds out.
Fill the 250 mL beaker with tap water so that it’s halfway filled and dip a section of the cut paper towel in so it’s completely wet.
This wetted paper towel section will be used to pre-germinate the seeds to ensure only viable seeds are planted in the soil at the beginning of the experiment.
Set out the paper towel section flat on the workspace, place all of the seeds around the centerline of the width of the paper towel, each approximately ½ cm away from each other. Fold the towel into fourths with the first fold being vertically and the second being horizontally.
Place the folded and wetted paper towel section into one of the 4 oz jars and close the jar.
This is so the paper towel stays wet and the seeds can germinate.
Using a piece of appropriately sized masking tape and a sharpie, label the jar “Neg”
Place the jar into the incubator and set the temperature to 23 ℃. If this is consistently around room temperature, just leave the jars out of the incubator.
Repeat steps 9-15 twice more to create another two sets of eighteen seeds for germination except, when doing step 7, label the jar “Pos” the second time, and “Exp” the third time.
Again, this is to ensure that there are at least nine viable germinated seeds for the experiment.
Using the stand on its back to stand it up, place one of the digital thermometers into the incubator.
This will help as it displays the temperature that the seeds are actually at.
Close and place the seed storage container back into the fridge.
Any seeds that don’t germinate after 18 days of being in the jar should be placed into the garbage (there may still be extra germinated seeds in the jar to ensure that all sections of the procedure have enough germinated seeds for planting). If there are no more seeds in the jar, throw away the paper towel and place the jar back into storage.
Temperature & Watering Section:
After the seeds have germinated and are planted (described in Seed Subsection steps 4-14 & Negative Control steps 11-20), using a 1 L graduated cylinder to measure, water each plant with 500 mL of tap water every three days.
Additionally, when watering plants for only the Positive Control plants, every two times the plants are watered (ie. 6 days) dissolve 10 g of the chemical fertilizer into every 500 mL of water before watering. This means that every six days, each plant from the Positive Control should receive 10 g of chemical fertilizer.
Refer to the 20-20-20 chemical fertilizer MSDS before handling.
Keep plants / the grow cart inside at all times as the temperature should always be between 21 ℃ and 27 ℃. If the temperature is ever outside the range, investigate why this may be the case and, if necessary, move the grow cart to a cooler location or open the flaps of the humidity tent.
Any temperature outside of these parameters could affect the growth of the plants and, therefore, the experiment.
Note that amaranth does not have any specific humidity requirements.
To ensure that the temperature is always in the range, every time the plants are watered, check the digital thermometer.
Glycerol Stock Section:
Put on all lab safety equipment, such as lab coat, goggles, and nitrile gloves
Take out all materials.
Using the black sharpie, label the ten 2 ml screw cap tubes “P.f. 1” - “P.f. 10” respectively so each tube is assigned a bacteria and unique number.
Using the bleach, tap water, and the 250 mL beaker, make a 5% bleach solution by adding 10 mL of bleach and 190 mL of tap water to the beaker. Do not let the bleach or bleach solution be touched by bare skin, inhaled, drank, or administered to the eyes. If any of this happens immediately rinse the affected area for several minutes or relocate to a place with fresh air.
Refer to the bleach MSDS before handling.
Plug the bunsen burner into the gas hose, turn on the gas, and use a lighter to light the flame. Make sure the burner isn’t immediately under anything and that the flame is blue and makes a triangle at its peak.
This is so all materials remain sterile as long as they’re close to the flame (due to the hot air rising and taking foreign bacteria with it).
Additionally, any non-flammable material (such as caps, lips of bottles, and inoculation loops) can be sterilized by running them through the flame.
Move all materials within a 6 inch radius of the burner.
Using glycerol, tap water, and a 15 ml container, make a 50-50 glycerol-water stock solution.
Using a 1 ml serological pipette, pipette 1 ml of 50-50 water-glycerol solution and 1 ml of P.f. liquid culture into the 2 ml screw cap tube labeled “P.f. 1”. Repeat this step for screw cap tubes labeled “P.f. 2” - “P.f. 10”.
Put all the tubes in a recycled plastic box (approximately 6” by 4” by 5”)
Put all the tubes in a freezer box and label the box “P.f. Glycerol Stocks” and the date it was made; store the solution in a -20 ℃ freezer.
Clean up all materials.
Repeat steps 1-12 except replace every instance of “P.f.” with “B.s.”.
Negative Arm:
Put on nitrile gloves and goggles (PPE).
Find a large enough work space so that 9 of the pots can fit on it in a 3x3 formation (48 cm x 45 cm space) and then place them on the workspace in that formation.
Using a ruler to measure, cut 9 sections of 8 cm long tape from the roll of masking tape.
After cutting each section of tape, so that the sections don’t lose their adhesiveness, hang them off of the side of the table.
Halfway down the pot, stick one 8 cm masking tape strip horizontally to the side of the pot to create a label. Repeat this for the 8 remaining pots and strips.
Using a black sharpie, label the tape on the 9 pots “Neg 1” - “Neg 9” respectively so each pot is assigned an experimental arm designation and unique number.
Using the scissors, open the Ocean Forest Potting Soil, the Ribbon Organics Compost, and the Biochar so that they can be closed by folding them and using a binder clip, but big enough so the garden scooper can transfer product out of the bags.
Using the garden scooper, fill each of the 9 pots up to the lip with a mix of 70% Potting Soil, 20% Compost, and 10% Biochar, making sure that all soil is flat and evenly distributed.
Store all pots labeled ‘Neg 1 - 9’ in their section of the grow cart so that all labels are facing forward.
Put away all excess materials (i.e. masking tape, sharpie, ruler, etc.), store PPE, and throw away any excess garbage.
This is a good place to pause as the next step involves waiting a large amount of time.
Three days after the seeds have been prepared for germination (steps 4-14 of the Seed Subsection), check the jars for germinated seeds (seeds that have opened and produced a root), and, if there are at least twenty-seven germinated seeds, move on to the next step. Otherwise, keep checking every day until there are twenty-seven.
Take out all germination jars in preparation to plant the germinated seeds.
Move all of the pots labeled ‘Neg 1 - 9’ back to the workstation and put PPE back on.
Using a ruler to measure, plant a germinated seed in the center of one of the pots 2.5 cm deep so that the roots face down. Repeat this process for the 8 remaining pots and germinated seeds.
Store any excess germinated seeds in the jar with the towel in the incubator to ensure that there are enough germinated seeds for other experimental sections.
Fill the 250 mL graduated cylinder up to the 100 mL mark with cool water from the sink and pour the water from the graduated cylinder into one of the pots evenly until all the soil is moist. Repeat this process for the other 8 pots.
Mark the labels of the pots with the date that the seeds were planted.
Bring all “Neg” labeled pots to the greenhouse area.
If the workstation rolls, roll the station over to the greenhouse to remove the hassle of carrying all of the pots. Otherwise, since the pots have handles, carry them over individually.
Place the pots on the greenhouse cart in the ‘Neg Section’ so that the labels are all facing forward.
This will also be referred to as the ‘Neg Section’ on data tables.
Go back to the workspace where the pots were originally filled with dirt and clean all materials used and return them to their respective places.
Refer to the plant care subsection of the care section for watering and temperature requirements.
Click on this link and this link and create a copy of both documents by selecting file and then ‘make a copy’ as they include templates for all data tables used in this experiment and places to record observations.
Make a Google Drive folder and title it ‘Plant Photo Storage’.
Every five days after planting, take a picture of each plant in the ‘Neg’ section of the grow cart using the phone. Store these photos in the ‘Plant Photo Storage’ Google Drive folder.
In the ‘Plant Photo Storage’ folder, label each photo taken with the corresponding label on the pot of the plant and with the days after planting (for example, ‘Neg 1’ day 5).
Using the photos, record observations of the ‘Neg’ section of the grow cart every five days in the Qualitative Data Table. These observations should include the color of the plant, number of leaves, lost leaves, spotting / disease, droopiness, height compared to plants in the same section, temperature of the grow cart, and more general observations about the health of the plant. Don’t record something like “the plants are doing well”.
At the ten day mark, use a ruler to measure the height of each of the plants (from the soil to the highest point) in cm. Record this data in its respective place in the ‘Negative control’ section of the Quantitative Data Table.
At the ten day mark, measure the girth of the stems of each plant in cm using the soft measuring tape at 1 cm above the base of the stalk. Record this data in its respective place in the ‘Negative control’ section of the Quantitative Data Table.
At the ten day mark, count the # of leaves on each of the plants in the ‘Neg’ section of the grow cart. Record this data in its respective place in the ‘Negative control’ section of the Quantitative Data Table.
Repeat steps 26-28 for the twenty, thirty, and forty day marks.
Place all extra materials back in storage, throw out all garbage, and put away PPE.
Positive Arm:
Repeat steps 1-30 of the negative control except for steps 21 & 22. Refer to steps 1 & 2 in the Temperature & Watering Subsection of the Plant Care Section for watering requirements. In all labeling, storage, and data recording steps replace ‘Neg’ with ‘Pos’.
(Non-condensed)
Put on nitrile gloves and goggles.
Find a large enough work space so that 9 of the pots can fit on it in a 3x3 formation (48 cm x 45 cm space) and then place them on the workspace in that formation.
Using a ruler to measure, cut 9 sections of 8 cm long tape from the roll of masking tape.
After cutting each section of tape, so that the sections don’t lose their adhesiveness, hang them off of the side of the table.
Halfway down the pot, stick one 8 cm masking tape strip horizontally to create a label. Do this for the 8 remaining pots and strips.
Using a black sharpie, label the tape on the 9 pots “Pos 1” - “Pos 9” respectively so each pot is assigned an experimental arm designation and unique number.
Using the scissors, open a bag of humus soil so that it can be closed by folding it and using a clip, but big enough so the garden scooper can transfer soil out of the container.
Using the garden scooper, transfer one bag (8 L) humus soil into each of the 9 pots making sure that all soil is flat and evenly distributed.
Take the “Seed Storage Container” out of the fridge and place it on the workstation.
Open the seed container and take eighteen seeds out.
Fill the beaker with tap water so that it’s halfway filled and dip a section of the 28 cm by 15 cm paper towel in so it’s completely wet.
This wetted paper towel section will be used to pre-germinate the seeds to ensure only viable seeds are planted in the soil at the beginning of the experiment.
Set out the paper towel section flat on the workspace, place nine of the eighteen seeds around the centerline of the width of the paper towel, each approximately ½ cm away from each other. Fold the towel into fourths with the first fold being vertically and the second being horizontally.
Place the folded and wetted paper towel section into one of the jars and close the jar.
This is so the paper towel stays wet and the seeds can germinate.
Place the jar into the incubator and set the temperature to 28℃.
Repeat steps 10-13 to create another set of nine seeds for germination.
Again, this is to ensure that there are at least nine viable germinated seeds for the experiment.
Close and place the seed storage container back into the fridge.
Store all pots labeled ‘Pos 1 - 9’ in their section of the grow cart so that all labels are facing forward.
Put away all excess materials (i.e. masking tape, sharpie, ruler, etc.), store PPE, and throw away any excess garbage.
This is a good place to pause as the next step involves waiting a large amount of time.
Wait three days, check the jars for germinated seeds (seeds that have opened and produced a root), and, if there are at least nine germinated seeds, move on to the next step. Otherwise, keep checking every day until there are nine.
Move all of the pots labeled ‘Pos 1 - 9’ back to the workstation and put PPE back on.
Using a ruler to measure, plant a germinated seed in the center of one of the pots 2.5 cm deep so that the roots face down. Repeat this process for the 8 remaining pots and germinated seeds.
Store any excess germinated seeds in the jar with the towel in the incubator to ensure that there are enough germinated seeds for other experimental sections.
Fill the 200 mL graduated cylinder up to the 100 mL mark with cool water from the sink and pour the water from the graduated cylinder into one of the pots evenly until all the soil is moist. Repeat this process for the other 8 pots.
Mark the labels of the pots with the date that the seeds were planted.
Bring all “Pos” labeled pots to the greenhouse area.
If the workstation rolls, roll the station over to the greenhouse to remove the hassle of carrying all of the pots. Otherwise, since the pots have handles, carry them over individually.
Place the pots on the greenhouse cart in the ‘Pos Section’ so that the labels are all facing forward.
This will also be referred to as the ‘Pos Section’ on data tables.
Go back to the workspace where the pots were originally filled with dirt and clean all materials used and return them to their respective places.
Refer to the Watering & Temperature Subsection of the Plant Care Subsection of the Care Section (steps 1-4) for watering, fertilizer, and temperature requirements. Remember that these are the Positive Control Section pots and should follow the specific requirements for them.
For the following steps, use the data tables that were copied in the Negative Control section to record data. Remember to use the ‘Positive Control’ tab in the Quantitative Data Table.
Every five days after planting, take a picture of each plant in the ‘Pos’ section of the grow cart using the phone. Store these photos in the ‘Plant Photo Storage’ Google Drive folder.
In the ‘Plant Photo Storage’ folder, label each photo taken with the corresponding label on the pot of the plant and with the days after planting (for example, ‘Pos 1’ day 5).
Using the photos, record observations of the ‘Pos’ section of the grow cart every five days in the Qualitative Data Table. These observations should include the color of the plant, number of leaves, lost leaves, spotting / disease, droopiness, height compared to plants in the same section, temperature of the grow cart, and more general observations about the health of the plant. Don’t record something like “the plants are doing well”.
At the ten day mark, use a ruler to measure the height of each of the plants (from the soil to the highest point) in cm. Record this data in its respective place in the ‘Positive control’ section of the Quantitative Data Table.
At the ten day mark, measure the girth of the stems of each plant in cm using the soft measuring tape at 1 cm above the base of the stalk. Record this data in its respective place in the ‘Positive control’ section of the Quantitative Data Table.
At the ten day mark, count the # of leaves on each of the plants in the ‘Pos’ section of the grow cart. Record this data in its respective place in the ‘Positive control’ section of the Quantitative Data Table.
Repeat steps 32-34 for the twenty, thirty, and forty day marks.
Place all extra materials back in storage, throw out all garbage, and put away PPE.
Experimental Arm:
Repeat steps 1-30 of the negative control except for steps 21 & 22 (before repeating, read through all steps in this arm). Refer to steps 1-4 in the Temperature & Watering Subsection of the Plant Care Section for watering requirements. In all labeling, storage, and data recording steps replace ‘Neg’ with ‘Exp’.
After step 17, complete the following steps before moving on.
Take all glycerol stocks filled with P. fluorescens out of the fridge and move them to the workstation. Wait 30 minutes for the cultures to warm up to room temperature before moving on to the next step.
Using the 50 mL serological pipette, transfer 0.5 mL from the screw cap labeled ‘P.f. 1’ into the pot labeled ‘P.f./B.s. 1’ directly on top of the germinated seed. Repeat this process for the other 8 pots.
Clean and wash out all screw caps with liquid culture inside using a 5% bleach solution as there will be excess liquid culture.
Refer to step 5 of the P.f./B.s. Culturing section for the process of making the bleach solution.
Repeat steps 2a - 2ci for glycerol stocks with B. subtilis.
Additionally, replace step 18 with the following: “Place the pots on the greenhouse cart in the ‘Exp Section’ so that the labels are all facing forward. Sequester the experimental pots as far away as possible from the control pots to prevent any possible cross contamination of bacteria.”.
(Non-condensed)
Put on nitrile gloves, goggles, and a lab coat (PPE).
Find a large enough work space so that 9 of the pots can fit on it in a 3x3 formation (48 cm x 45 cm space) and then place them on the workspace in that formation.
Using a ruler to measure, cut 9 sections of 8 cm long tape from the roll of masking tape.
After cutting each section of tape, so that the sections don’t lose their adhesiveness, hang them off of the side of the table.
Halfway down the pot, stick one 8 cm masking tape strip horizontally to the side of the pot to create a label. Repeat this for the 8 remaining pots and strips.
Using a black sharpie, label the tape on the 9 pots “Exp 1” - “Exp 9” respectively so each pot is assigned an experimental arm designation and unique number.
Using the scissors, open a bag of humus soil so that it can be closed by folding it and using a clip, but big enough so the garden scooper can transfer soil out of the container.
Using the garden scooper, transfer one bag (8 L) humus soil into each of the 9 pots making sure that all soil is flat and evenly distributed.
Take the “Seed Storage Container” out of the fridge and place it on the workstation.
Open the seed container and take eighteen seeds out.
Fill the beaker with tap water so that it’s halfway filled and dip a section of the 28 cm by 15 cm paper towel in so it’s completely wet.
This wetted paper towel section will be used to pre-germinate the seeds to ensure only viable seeds are planted in the soil at the beginning of the experiment.
Set out the paper towel section flat on the workspace, place nine of the eighteen seeds around the centerline of the width of the paper towel, each approximately ½ cm away from each other. Fold the towel into fourths with the first fold being vertically and the second being horizontally.
Place the folded and wetted paper towel section into one of the jars and close the jar.
This is so the paper towel stays wet and the seeds can germinate.
Place the jar into the incubator and set the temperature to 28 ℃.
Repeat steps 10-13 to create another set of nine seeds for germination.
Again, this is to ensure that there are at least nine viable germinated seeds for the experiment.
Close and place the seed storage container back into the fridge.
Store all pots labeled ‘Exp 1 - 9’ in their section of the grow cart so that all labels are facing forward.
Put away all excess materials (i.e. masking tape, sharpie, ruler, etc.), store PPE, and throw away any excess garbage.
This is a good place to pause as the next step involves waiting a large amount of time.
Wait three days, check the jars for germinated seeds (seeds that have opened and produced a root), and, if there are at least nine germinated seeds, move on to the next step. Otherwise, keep checking every day until there are nine.
Move all of the pots labeled ‘Exp 1 - 9’ back to the workstation and put PPE back on.
Using a ruler to measure, plant a germinated seed in the center of one of the pots 2.5 cm deep so that the roots face down. Repeat this process for the 8 remaining pots and germinated seeds.
Store any excess germinated seeds in the jar with the towel in the incubator to ensure that there are enough germinated seeds for other experimental sections.
Fill the 200 mL graduated cylinder up to the 100 mL mark with cool water from the sink and pour the water from the graduated cylinder into one of the pots evenly until all the soil is moist. Repeat this process for the other 8 pots.
Mark the labels of the pots with the date that the seeds were planted.
Take all tubes filled with the P. fluorescens liquid culture out of the fridge and move them to the workstation. Wait 30 minutes for the cultures to warm up to room temperature before moving on to the next step.
Using the 50 mL serological pipette, transfer 25 mL from the tube labeled ‘Exp 1’ into the pot labeled ‘Exp 1’ directly on top of the germinated seed. Repeat this process for the other 8 tubes and pots.
Clean and wash out all tubes with liquid culture inside using a 5% bleach solution as there will be excess liquid culture.
Refer to step 5 of the P.f./B.s. Culturing section for the process of making the bleach solution.
Bring all “Exp” labeled pots to the greenhouse area.
If the workstation rolls, roll the station over to the greenhouse to remove the hassle of carrying all of the pots. Otherwise, since the pots have handles, carry them over individually.
Place the pots on the greenhouse cart in the ‘Exp Section’ so that the labels are all facing forward. Sequester the experimental pots as far away as possible from the control pots to prevent any possible cross contamination of bacteria.
This will also be referred to as the ‘Exp Section’ on data tables.
Go back to the workspace where the pots were originally filled with dirt and clean all materials used and return them to their respective places.
Refer to steps 1-4 of the Watering & Temperature Subsection of the Plant Sare subsection of the Care Section for watering and temperature requirements.
For the following steps, use the data tables that were copied in the Negative Control section to record data. Remember to use the ‘Experimental Arm’ tab in the Quantitative Data Table.
In the ‘Plant Photo Storage’ folder, label each photo taken with the corresponding label on the pot of the plant and with the days after planting (for example, ‘Exp 1’ day 5).
Using the photos, record observations of the ‘Exp’ section of the grow cart every five days in the Qualitative Data Table. These observations should include the color of the plant, number of leaves, lost leaves, spotting / disease, droopiness, height compared to plants in the same section, temperature of the grow cart, and more general observations about the health of the plant. Don’t record something like “the plants are doing well”.
At the ten day mark, use a ruler to measure the height of each of the plants (from the soil to the highest point) in cm. Record this data in its respective place in the ‘Experimental control’ section of the Quantitative Data Table.
At the ten day mark, measure the girth of the stems of each plant in cm using the soft measuring tape at 1 cm above the base of the stalk. Record this data in its respective place in the ‘Experimental control’ section of the Quantitative Data Table.
At the ten day mark, count the # of leaves on each of the plants in the ‘Exp’ section of the grow cart. Record this data in its respective place in the ‘Experimental control’ section of the Quantitative Data Table.
Repeat steps 34-36 for the twenty, thirty, and forty day marks.
Place all extra materials back in storage, throw out all garbage, and put away PPE.
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