Overview

Our project focused on two different pathways that can be used to increase phosphate uptake. One of them used glucose dehydrogenase to remove the heavy metals in mineral phosphates, thereby increasing the ability of the plants to uptake the phosphate. The other amplified expression of the pstSCAB operon, which is already present in Rhizobacteria, with the intent of increasing the number of ion channels in the membrane of the Rhizobium. All of the coding sequences for both of these pathways will be included below, as well as the promoters, ribosome binding sites, and terminators used in the creation of the composite parts.

Pqq/gcd Pathway

Basic Parts

Composite Parts

pstSCAB Pathway

Basic Parts

Composite Parts

Other Parts

In addition to these coding sequences, we also cataloged 3 different promoters and 2 primer sequences

Testing media growth rates (July 2023)

For this experiment, we explored the growth rates of Rhizobium tropici in three different media: ATCC medium 111, high calcium yeast mannitol, and low calcium yeast mannitol.

Introduction: ATCC Medium 111, also known as Rhizobium X Medium, is a type of soil extract medium used in microbiology. It's typically used for the growth and propagation of certain types of bacteria, such as Rhizobium tropici. The specific composition of the medium can support the survival, proliferation, and biological function of these organisms. We choose to test this medium because our bacteria dwells in the rhizosphere, and we believed this media would act as a close representation to the real environment.

Results: A t-test of two samples for the means between the low Ca concentration yeast mannitol and high Ca concentration yeast mannitol yielded a p-value of 0.11. Therefore, since there was no statistical difference in growth; it was decided to use the low Ca yeast mannitol media since a low Ca was preferred for increased accuracy of measurements and lower cost and there was no evidence to suggest that the yeast mannitol with higher concentrations of Ca would yield higher rhizobium ODs.

Procedures for making media:

ATCC Medium 111 (Rhizobium X Medium):

Introduction: ATCC Medium 111, also known as Rhizobium X Medium, is a type of soil extract medium used in microbiology. It's typically used for the growth and propagation of certain types of bacteria, such as Rhizobium tropici. The specific composition of the medium can support the survival, proliferation, and biological function of these organisms. We choose to test this medium because our bacteria dwells in the rhizosphere, and we believed this media would act as a close representation to the real environment.

Materials:

• Yeast extract 1.0g
• Mannitol 10.0g
• Agar 15.0g
• Distilled water 800.0mL
• Soil extract 200.0mL
• African violet soil 77.0g
• Na2CO3 0.2g
• Distilled water 200.0mL

Procedure:

• Combine all materials
• Adjust pH to 7.2 and autoclave with 15lbs. pressure at 121 C for one hour to sterilize
• Filter medium through paper to remove soil before using

The final product should be translucent and tan in color. There may be a slight precipitate. It is recommended to store the medium at a temperature between 2-8 C.

Yeast Mannitol:

Introduction: Yeast Mannitol Broth is used for the cultivation and enumeration of soil microorganisms like Rhizobium species. Yeast extract provides a source of readily available amino acids, vitamin B complex, and accessory growth factors for Rhizobia. Mannitol is the carbohydrate source. The pH of the medium is adjusted to 6.8±0.2 at 25 C. In addition to Rhizobium, this medium can also be used for other symbiotic nitrogen-fixing soil microorganisms.

Materials:

• Yeast extract 1.0g
• Mannitol 10.0g
• Dipotassium phosphate 0.5g
• Magnesium sulfate 0.2g
• Sodium chloride 0.1g
• Calcium carbonate (use 1.0g for high calcium media and 15mg for low calcium media)
• Distilled water 1.0L

Procedure:

• Suspend 12.8 g of Yeast Mannitol Broth in 1000 ml of distilled water.
• Boil to dissolve the medium completely
• Sterilize by autoclaving at 15 lb. pressure at 121 C for 15 minutes

Transformation Test (July 2023):

For this experiment, we tested whether our protocol for plasmid insertion was effective. Our empty plasmid contained Kanamycin antibiotic resistance and lacZ genes with the inducible lac promoter. We used blue-white screening to confirm integration.

X-gal, or 5-bromo-4-chloro-3-indolyl β-D-galactopyranoside, is a chromogenic substrate for the enzyme β-galactosidase. This means it changes color when it is cleaved by the enzyme. In molecular biology, X-gal is used in a technique known as blue-white screening, which is a rapid and efficient method for the identification of recombinant bacteria.

A plasmid vector is engineered to contain a segment of the lacZ gene, which codes for β-galactosidase. This plasmid is taken up by R. tropici cells through a transformation of electro-competent cells(https://2014.igem.org/Team:Hannover/Protocols/Rhizobium_Electroporation). When X-gal is added to the growth medium, it is cleaved by any β-galactosidase that is present, producing an insoluble blue pigment. If a foreign DNA sequence is inserted into the lacZ gene on the plasmid (where we designed our multiple cloning site to be located), it disrupts the gene and prevents it from producing β-galactosidase.

Therefore, colonies of bacteria that have taken up the plasmid but have not had foreign DNA inserted into the lacZ gene (non-recombinant) will turn blue. In contrast, colonies that have taken up the plasmid and have had foreign DNA inserted into the lacZ gene (recombinant) will remain white because they cannot cleave X-gal. This allows for easy visual identification of recombinant bacteria.

Results: We were able to confirm the insertion of an empty plasmid with blue-white screening; our colonies were blue.

Ligation (August 2023):

Introduction: Ligation is a process in molecular biology that involves the joining of two nucleic acid fragments through the action of an enzyme. It is an essential laboratory procedure in the molecular cloning of DNA, whereby DNA fragments are joined to create recombinant DNA molecules. This process can occur when a foreign DNA fragment is inserted into a plasmid. For our project, we needed to join two gcd gene fragments with a PqqC gene fragment.

We performed ligation using T4 DNA ligase. This enzyme is broadly used in vitro as a molecular biology research tool due to its capability of joining both sticky and blunt DNA ends. The ends of DNA fragments are joined by the formation of phosphodiester bonds between the 3’-hydroxyl of one DNA terminus with the 5’-phosphoryl of another. This process is catalyzed by an enzyme called DNA ligase.

Results: Successful ligation is proved by PCR performed after gene insertion.

Protocol:

Set up the following reaction in a microcentrifuge tube on ice. (T4 DNA Ligase should be added last. Note that the table shows a ligation using a molar ratio of 1:3 vector to insert for the indicated DNA sizes.)

Procedure:

• Gently mix the reaction by pipetting up and down and microfuge briefly.
• For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes.
• For blunt ends or single base overhangs, incubate at 16°C overnight or room temperature for 2 hours.
• Heat inactivate at 65 C for 10 minutes.
• Chill on ice and transform 1-5 μl of the reaction into 50 μl competent cells.

Plasmid Gene Insertion (August-September):

Introduction: The NEBuilder® HiFi DNA Assembly Cloning Kit is a tool developed to improve the efficiency and accuracy of DNA assembly. It allows for seamless assembly of multiple DNA fragments, regardless of fragment length or end compatibility. It can be used to assemble either single-stranded oligonucleotides or different sizes of DNA fragments with varied overlaps (15–30 bp). The kit is useful for the synthetic biology community, as well as those interested in one-step cloning of multiple fragments due to its ease of use, flexibility, and simple master-mix format. It can join DNA fragments together more efficiently, even with larger fragments or low DNA inputs. The kit can be used in successive rounds of assembly, as it removes 5´ and 3´ end mismatches.

Preparation Protocol:

• pstSCAB contains 4 genes (pstS, pstC, pstA, & pstB), add 1μL of each
• PqqC + gcd contains 3 genes (PqqC, gcd 1, & gcd2), add 1μL of each
• 300 ng of back bone (2 μL)
• 10μL of master mix
• add water to end of with 20 μL total
• Incubate for 1 hour @ 50 C

Assembly Protocol:

• Thaw and vortex master mix thoroughly and keep on ice
• Incubate samples in a thermocycler at 50 C for 15 minutes when 2 or 3 fragments are being assembled or 60 minutes when 4 - 6 fragments are being assembled. Following incubation, store samples on the ice at - 20 C for subsequent transformation
• Transform competent cells with 2 μL of the assembly reaction, using the following the transformation protocol

Chemically Competent Cells Transformation Protocol:

• Thaw chemically competent cells on ice
• Add 2 μL of the chilled assembled product to the competent cells. Mix gently by pipetting up and down by flicking the tube 4 - 5 times. Do not vortex
• Place the mixture on ice for 30 minutes. Do not mix.
• Heat shock at 42 C for 30 seconds. Do not mix.
• Transfer tubes to the ice for 2 minutes
• Add 950 μL of room-temperature SOC media to the tube
• Incubate the tube at 37 C for 60 minutes. Shake vigorously (250 rpm) or rotate
• Warm selection plates to 37 C
• Spread 100 μL of the cells onto the selection plates. Use Amp plates for the positive control sample
• Incubate overnight at 37 C

Gene Confirmation with Polymerase Chain Reaction (PCR) and Gel Electrophoresis (September 2023):

For this experiment, we tested whether our genes were integrated into our plasmids. We performed a colony PCR with Q5 and a gel electrophoresis. Colony PCR is a high-throughput method for determining the presence or absence of insert DNA in plasmid constructs. Specifically, we wanted to confirm our genes by confirming the insert length (from the colony PCR) with gel electrophoresis.

Results: As seen in the images below, we were able to confirm the length of insertion fragments. We expected a fragment length of around 3700 base pairs for pstSCAB and a fragment length of around 3200 for PqqC + gcd.

Colony PCR Protocol:

Materials:

• 10uM primer VF2
• 10uM primer VR
• 2X Q5 HotStart Master Mix
• Distilled water
• Agar plate with bacterial colonies
• PCR tubes
• Toothpicks

Collecting Bacterial Cultures + Reaction Setup:

Be sure that you save each bacterial colony so that if the PCR shows what you want, you can go back to the original bacterial cells. You can do this by dipping the toothpick in step 5 below into liquid media or using to swab an agar plate before adding the DNA into the PCR tube.

• Add 1.3 ul of primer VF2 to PCR tube
• Add 1.3 ul of primer VR to PCR tube
• Add 12.5 ul 2x Master Mix to PCR tube
• Add 10 ul of water to PCR tube
• Using toothpick, take 1 colony of the bacteria from the agar plate and add it to the PCR tube (step 5)
• Put PCR tube into thermocycler and run using the following parameters:

PCR Cycle:

• Initial denaturation @ 98 C for 30 seconds

25-30 Cycles of the following:

• 98 C for 5-10 seconds
• 50-72 C for 10-30 seconds (variable depending on primers used; select temperature based on primer annealing)
• 72 C for 20-30 seconds per kilobase
• Final extension @ 72 C for 2 minutes
• Hold @ 4-10 C

Gel Electrophoresis Protocol:

Preparing the gel:

• Weigh out 0.5g of the agarose on a piece of weigh paper. Transfer to an Erlenmeyer flask. Add 50 ml of 1X TAE.
• Place the flask in the microwave for 30 seconds to 1 minute. Heat until the agarose and buffer mixture is fully dissolved and remove with cooking mittens. Periodically swirl mixture to ensure the mixture is melting evenly.
• Remove the flask of clear agarose and allow it to cool. This will take about 10 minutes. If you are able to grasp the flask comfortably it is cooled enough to pour.
• While the agarose is cooling, place the gel tray into the gel box and add blocker and a comb.
• When the agarose is cooler, add 2 ul of Apex Safe DNA Gel Stain to the melted agarose.
• Swirl the agarose to incorporate the dye and pour the agarose into the gel tray—your gel should be about 1 cm thick.
• Allow at least 20 minutes for the gel to solidify. Once solid, carefully remove the comb and blockers, and place the solidified gel (still on the tray) oriented on the negative side.
• Add enough 1X TAE buffer to completely cover the gel by 1 cm.

Preparing the Sample:

On a piece of parafilm, spot 1 ul of 6X DNA loading dye with a p20 pipette. You should have one spot for each of your PCR products. Wait until your gel has started to solidify before beginning the step since the spots will dry out after about 10 minutes. You can also do this step in a tube.

• Add 5ul of water to each spot of dye.
• Add 5ul of the PCR product.

Running a Gel:

• Into the first lane of the gel load 5ul of the 2-log DNA ladder.
• Into lane 2-5 load 10ul of each of your PCR products including the water and dye.
• Place gel lid with electrodes on the gel box.
• Set the voltage to 100 volts. You should see bubbles coming up from the wire electrode.
• After a few minutes, check to make sure everything is going smoothly. Run the gel for approximately 30 minutes or until the dye is 2/3 the way down the gel.
• Take a picture of the gel.
• Use the DNA ladder to approximate the size of each of your DNA bands. Make sure you check that your DNA band is the correct size.

Phosphate Uptake Assays (August-September):

For this experiment, we wanted to confirm whether our engineered Rhizobium would more effectively uptake an inorganic phosphate. During our initial control tests, we found that our strain of R. tropici was unable to uptake inorganic phosphate, making it a negative control. Therefore, we used Pseudomonas fluorescens as a positive control because it was present in our lab and has been shown to uptake inorganic phosphate in numerous literature.

To do this, we made agar plates with a modified low calcium yeast mannitol that replaced the dipotassium phosphate with tricalcium phosphate. Next, we put bacterial spot dilutions and let them incubate for 7 days. After the incubation period, we observed the “halo” – a clearing made around the bacterial spot – and compared the area.

We also performed liquid assays to account for the high motility of our positive control, P. fluorescens. We made 22 liquid media samples with the protocol for the tricalcium phosphate yeast mannitol. We also made 9 agar plates with tricalcium phosphate media. All bacterial solutions were 10uL in 5mL of tricalcium phosphate yeast mannitol liquid media. Samples 1-6 did not contain kanamycin or IPTG. Samples 7-22 contained 25uL kanamycin and 50uL IPTG per sample. Plates 1 and 2 did not contain kanamycin. 1x = 200uL of bacterial solution spun down with 190uL of supernatant removed. 2x = 400uL of bacterial solution spun down with 385uL of supernatant removed. P. fluorescens 1x = 500uL of bacterial solution spun down with 490uL of supernatant removed. P. fluorescens 2x = 1000uL of bacterial solution spun down with 985uL of supernatant removed.

Protocol:

Materials:

• Yeast extract 0.5g
• Mannitol 5.0g
• Magnesium sulfate 0.1g
• Sodium chloride 0.05g
• Calcium carbonate 0.005g
• Tricalcium phosphate 2.5g
• Agar 7.5g
• Distilled water 500mL

Procedure:

• Swirl to combine all materials in a 1L glass
• Sterilize by autoclaving at 15lb pressure at 121 C for 15 minutes
• Let it cool to 50 C (should be able to touch with bare hand without burning)
• Add 50ng/uL kanamycin; swirl to combine
• Pour plates

Testing Phosphate Assays:

• Spread 100uL 100mM IPTG before adding bacterial spots
• Let plates dry under laminar flow hood for about 1 hour
• Wrap with parafilm to ensure moisture is not lost and incubate at 32 C for 7 to 10 days

Phosphate Solubilization Assays (September-October):

The colorimetric phosphate assay is a method used to measure the concentration of phosphate in a sample.

In a dilute orthophosphate solution, ammonium molybdate reacts under acid conditions to form the heteropoly acid, molybdophosphoric acid. In the presence of vanadium, yellow vanadomolybdo-phosphoric acid is formed. The intensity of the yellow color is proportional to the phosphate concentration. The Phosphate Colorimetric Assay Kit provides a simple and direct procedure for measuring phosphate levels (ranging from 1–5 nmole/well) in a variety of samples. Phosphate reacts with a chromogenic complex, which results in a colorimetric (650 nm) product proportional to the amount of phosphate present.

The sample is mixed with the vanadate-molybdate reagent and the absorbance is measured. The phosphate concentration in the sample can then be determined by comparing its absorbance with that of the calibration standards.

Procedure

• All samples and standards should be run in duplicate.

Phosphate Standards for Colorimetric Detection

• Dilute 10 mL of the 10 mM Phosphate Standard with 990 mL of water to prepare a 0.1 mM Phosphate Standard Solution.
• Add 0, 10, 20, 30, 40, and 50 mL of the 0.1 mM Phosphate Standard Solution into a 96 well plate, generating 0 (blank), 1, 2, 3, 4, and 5 nmole/well standards.
• Add water to each well to bring the volume to 200 mL.

Sample Preparation

Samples can be measured directly.

• Add 1–200 mL of sample to wells.
• Bring samples to a final volume of 200 mL with water.

Note: For unknown samples, it is suggested to test several sample dilutions to ensure the readings are within the linear range of the standard curve.

Assay Reaction

• Add 30 mL of the Phosphate Reagent to each of the wells.
• Mix well using a horizontal shaker or by pipetting, and incubate the reaction for 30 minutes at room temperature.
• Cover the plate and protect from light during the incubation.
• Measure the absorbance at 650 nm (A650).

Note: When using 1.0 mL cuvettes, increase the volume of all reaction components 5-fold. The 1 mL total reaction volume will contain 1–500 mL of sample, 150 mL of Phosphate Reagent, and bring to a final volume of 1 mL with water.

• Incubate at room temperature for 30 minutes.
• Measure the absorbance at 650 nm (A650).

Protein Gel (October):

The main purpose of a protein gel, also known as protein gel electrophoresis, is to separate proteins based on their size. We ran a protein gel to confirm the synthesis of our various parts. The fragment lengths we were looking for are:

gcd: GDH enzyme- 86,747 mass (Da)

PqqC: Pqq synthase- 47,425 mass (Da)

pstS: pstS Pi binding protein- 37,024 mass (Da)

pstC: pstC membrane transport protein- 34,121 mass (Da)

pstA: pstA membrane transport protein- 32,322 mass (Da)

pstB: pstB ATP-binding protein- 29,027 mass (Da)

Results: Our first protein gel was unsuccessful as the bars were unclear and under 25000Da.

Protocol:

Cell Lysis:

• Discard the medium in culture dishes with cells and wash the cells using ice-cold Phosphate-buffered saline (PBS).
• Discard the PBS, add ice-cold lysis buffer.
• Scrape the cells using a cold plastic cell scraper.
• Collect the cells in microcentrifuge tubes.
• Agitate the contents in microcentrifuge tubes for 30 min at 4 °C.
• Centrifuge the tubes at 16,000 x g for 20 min at 4 °C.
• Collect the supernatant in a fresh tube and place on ice. Discard the pellet.

Sample Preparation:

• To a volume of protein sample (cell lysate), add an equal volume of loading buffer.
• Boil the above mixture at 95 °C for 5 min.
• Centrifuge at 16,000 x g for 5 min.
• These samples can be stored at -20 °C or may be used to proceed with gel electrophoresis.

Gel Electrophoresis:

• Load your samples into wells created in the stacking gel.
• Apply an electric field across the gel. The proteins will migrate through the gel based on their charge and size.
• Once electrophoresis is complete, stain the gel to visualize the proteins. This can be done using various methods such as Coomassie dye staining or silver staining.
• Remove excess dye from the gel matrix background.

Results: Our second protein gel showed promising results for gcd protein synthesis. We expected 87 kDa and saw a corresponding band for one of our PqqC+gcd samples.

Plant Experiments:

Our ultimate goal is to measure the micro symbiosis between our Rhizobium and our various plant species (Common Bean, Soybean, Showy Pink Trefoil, American Senna.). We need to do this, because we want to know if we can implement the project using native plants as our micro symbionts.

We are also trying to measure if our Rhizobium is actually beneficial for the plant's growth/if it experiences phosphate poisoning. If we find negligible differences in growth overtime, if we see no differences, then we will know that either the Rhizobium never took root (hehe) or that the Rhizobium doesn’t help the plant that much. We need to do this in order to see if our “built in killswitch” theory holds true.

The first thing we do is acquire a wet paper towel, and drape it on top of a ziploc bag. Then, we place the chosen amount of seeds on one half of the paper towel. Fold in half, and place the paper towel inside of the ziplock bag. We do this so that we can track germination. Rhizobium has a limited window of 3 days after germination for inoculations to be successful.

Once germinated, transfer GERMINATED SEEDS EXCLUSIVELY into a small planter. Inoculate a portion of the seeds in dirt with Rhizobium.(One portion will be a control). Once the plant outgrows the planter, gently transfer to a pot. We do this to prevent our plant from getting root bound. Let the plant grow ⅔ of the way until maturity, and then that is when you begin nodulation checks.

In the beginning, we planned a series of experiments to test the optimum conditions for our Rhizobium to survive. This included assessing the best microsymbiont, and assessing what the best age for inoculating is.

However, we have unfortunately been pressured to annul the age experiments. Our first Rhizobium check proved fruitless aside from a single set of modules on a showy pink trefoil.

As a result, we want to maximize our chances with inoculations, so we have been removing innoculatory controls from experiments. Since we have measured all previous growth rates, but they proved to be invalid, we have deemed them to be considered as controls.

For the past week or two we have been studying a hypothesis that it may be temperature that caused our lack of success. As a result, we are now incubating our plants at 29° C. We will then contrast the growth rates with our previous rates.

Important dates:

• Began plant experiments 7/15/2023
• First inoculation 7/27/2023
• First nodulation check 9/2/2023
• First nodules found 9/9/2023
• Temperature Hypothesis Transfer 9/23/2023
• Final Nodule Check 10/7/2023