The phosphoinositide (PI) pathway is involved in sensing changes in gravity, osmotic stress, and light-mediated signaling in plants. The nature of the regulation of the pathway is currently unknown, but is hypothesized to involve the binding of proteins to the key enzyme phosphatidylinositol 4 phosphate 5-kinase (PIPK). To test this hypothesis, I have been attempting to generate plant cells expressing tandem affinity purification (TAP)-tagged forms of the PIPK in order to recover interacting proteins for further analysis. Three approaches were used to achieve this goal: vector-mediated transformation of Arabidopsis cell liquid cultures through co-cultivation with the bacterium, Agrobacterium tumefaciens; a similar method involving callus growth of co-cultivated cell cultures; and a floral dip method involving the submersion of whole Arabidopsis flowers into a solution of A. tumefaciens.
Conditions were optimized for the liquid culture transformation; however, this potentially rapid transformation method has yet to yield transformants. Callus growth experiments have produced putative transformed calli and the plants used in the floral dip procedure have begun producing seeds. In the future, I will check for the presence of the TAP-tag mRNA and tagged protein. Once the presence of the protein is confirmed, I will purify the TAPtagged PIPK and optimize conditions for the recovery of interacting proteins.
Plants are sessile organisms that rely on a complex system of signaling pathways to sense and respond to changes in their environment. One pathway, which is important in sensing changes in osmotic stress (as reviewed in Munnik and Vermeer. 2010) and gravity sensing
in plants is the phosphoinositide (PI) pathway. The PI pathway involves the hydrolysis of phosphatidylinositol(4,5)bisphosphate (PtdIns(4,5)P
) by phospholipase C to produce the second messenger inositol trisphosphate (InsP
) (Figure 1). The PI pathway is found in all eukaryotic organisms studied thus far; however, the regulation of the pathway appears to be quite different in plants. Plant cells, in general produce much less PtdIns(4,5)P
2 than animal cells and, as a consequence, less InsP
is synthesized by the phosphorylation of phosphatidylinositol 4 phosphate (PtdIns4P) by an enzyme called PtdIns4P 5-kinase (PIPK). Scientists proposed that PIPK activity was the flux limiting step in the plant PI pathway and demonstrated that plant PIPKs were much less active than the animal
PIPKs in vivo.
The question that has eluded plant scientists pertains to why the plant PIPK is so much less active.
One clue is that the major isoforms of plant PIPKs have an additional N-terminal domain not found in the animal proteins. Specifically, the N-terminal domain of the Arabidopsis PIPK1 contains 7, 23-amino acid membrane occupational and recognition nexus (MORN) motifs.
The MORN motifs are found in other proteins that are involved in membrane fission and fusion; however, of all eukaryotic organisms reported thus far, only the plants have PIPKs with this unusual N-terminal MORN domain. (Figure 2)
When the MORN domain is removed, the specific activity of the plant enzyme, PIPK1, increased 4-6 fold, making it more similar to the animal PIPK.
The work previously done in our lab by Im, et al. suggests the N-terminal MORN domain binds selective plant proteins that are critical for controlling the activity of the plant PIPKs in vivo. In order to
test this hypothesis, we proposed to compare the proteins that bind the MORN domain of Arabidopsis
PIPK1 and the full-length protein, AtPIPK1. In addition, we proposed to take advantage of the fact that
Arabidopsis PIPK activity increases in response to hyperosmotic stress
in order to compare the binding proteins in stressed and non-stressed plants. Our assumption was that different key regulator proteins would bind in the stressed and non-stressed conditions.
In order to characterize these interacting proteins, we attempted to transform the model plant
Arabidopsis thaliana with a form of the AtPIPK1 gene tagged with a tandem affinity purification tag
(TAP-tag). Using dual-affinity binding purification (as diagrammed in Figures 3A and 3B), we intended to isolate and analyze the proteins which bind to both the full-length AtPIPK1 (FL) and the MORN domain specifically. Our plans were to transform both cells grown in suspension culture and whole plants. Once we obtained transformed cells, we would purify the AtPIPK1 and MORN interacting proteins by using the TAP-tag method.
Despite the use of several protocols and variations of procedures, viable transformants have yet to be produced and the initial step of the project, budgeted to take only two months, has become the project itself. Throughout the spring and summer months, we have been optimizing conditions of transformation in order to create transgenic A. thaliana plants carrying the genes for the TAP-FL and
TAP-MORN constructs. The optimized conditions and an analysis of the putative transforms are presented in this report.
Floral dip involves the direct inoculation of an unopened flower bud into a bacteria culture. The procedure followed was from A. Wiktorek-Smagur, 2009.
Specifically, wild-type plants were grown in growth chambers, a process which takes approximately 14-20 days from seed. A. tumefaciens containing the desired plasmid had been kept frozen at -80 o
C. The bacteria were spread on selectable YEP plates
(20ug/mL rifampicin, 300ug/mL spectinomycin) for two days. Colonies were picked and inoculated in
3mL liquid YEP media containing the same concentration of antibiotics and grown at 28 o
C for 2-3 days in an incubator shaker. The 3mL cultures were used to inoculate larger 600mL cultures and grown for an additional day. When an OD
of 0.8 was reached, the bacterial culture was centrifuged and the supernatant was removed. The pellet of cells was then resuspended in ½ MS with a 5% sucrose solution with added acetosyringone, which aids in the binding of the bacteria, and Silwet L-77, a surfactant. The floral buds of the plant were dipped into the mixture for up to two minutes. The buds were removed and left in the dark for 48 hours before returning them to normal growth conditions of 24hr light at
C. Siliques containing seeds were collected once they matured on the plant and kept dry in the lab.
To screen for transformed plants, the seeds were surface sterilized and plated on Murashige and Skoog
(MS) media + 100ug/mL gentamycin, the selectable marker. Once seedlings began to grow true leaves or root into the media, they were transferred to soil and grown in a short-day cycle growth chamber until enough leaf tissue has been produced to analyze for the presence of the Tap-tag transcript.
Growth rates of liquid Arabidopsis cell cultures were determined using six identical pairs of aliquots of
Arabidopsis cells. 1 or 2mL of cells were inoculated into each flask of 25mL Gamborg’s B5 vitamin media.
Cells were grown in a temperature-controlled shaker at 133 rpm and 21 o
C. Cells were collected from one pair of flasks after 48 hours via gravity filtration and their mass recorded. This was repeated every
24 hours thereafter until 6 trials had been completed.
The protocol for transformation of cells grown in suspension culture was based primarily on the one published by Salinas and Sanchez-Serrano
, although a number of papers were used to optimize conditions.
Arabidopsis cell cultures were grown in Gamborg’s B5 media in a temperature-controlled shaker at 133 rpm and 21 o
C. The cells should be optimal for transformation just before entering their
logarithmic growth phase, 2 to 3 days after introduction to new media. Bacteria containing the desired
Tap-tag were grown as described above; however, a much lower innoculum was used (100μL of bacteria culture per 10mL of plant cell culture). The bacteria were grown to an OD
600 of 1.0 before being centrifuged, washed, and resuspended in Gamborg’s B5. The Arabidopsis cells were then co-cultivated with 100uL of the diluted A. tumefaciens. The bacteria and plant cells were co-cultivated for 48 hours in the dark, during which time the A. tumefaciens bound to the Arabidopsis cell surface and insert their genes into the plant cell genome. Aliquots of the cells were removed under sterile conditions and viewed under a light microscope to confirm the presence and binding of the A. tumefaciens to the plant cell surfaces (Figure 4). After co-cultivating, the culture was washed via centrifugation to remove unbound bacteria and the remaining plant cells were grown on agar plates with the selectable antibiotic present. The cells which were successfully transformed should grow into a callus that could have been resuspended in fresh medium to initiate the new, transgenic culture.
Synthesis of a Fluorscein Isothiocyanate-Glutamine (FITC-Gln) Conjugate
To synthesize FITC-Gln, 13mg of free glutamine was reacted with 5mg of fluorescein isothiocyante (FITC) in 0.1M NaHCO
, pH8.0 to form a fluorescently labeled glutamine molecule (FITC-Gln). As the FITC formed bonds with the amine group of the glutamine, an expected color change from orange to yellow was observed. Free fluorescein is green and was used as a control. 7-14 day old WT and Hs9-7 seedlings were exposed to FITC-glutamine by incubation in MS media containing minimal amounts (roughly 0.1% v/v) of the fluorescent molecules. The seedlings were rinsed in medium without fluorescent compounds and intracellular fluorescence was monitored using a Nikon Optiphot fluorescent microscope with a U excitation filter and 514 emission filter. Fluorescence was recorded in transgenic and non-transgenic
(WT seedlings) using a Leica camera after four hours of incubation in the FITC-Gln solution.
Thousands of seeds were collected from the putative transformed plants. Nothing survived when the seeds were screened on the selection medium, MS media + 100μg/mL gentamycin. This meant that either nothing had been transformed or that the gentamycin concentration used was too strong or the growth conditions were not appropriate such that all the transformed seedlings were killed. To test these hypotheses, we first varied the gentamycin concentration by trying lower concentrations of
50ug/mL and 75ug/mL. In addition, the plants were moved from a 24-hour light exposure to a short day cycle to reduce the stress that might result from the longer light period. Neither the changes in gentamycin concentration nor the light regime improved survival of the putative transformants. Because we might have generated transgenics that were sensitive to the selection medium, we began transferring the most successful-looking putative transformants which had begun growing true leaves to
MS media without antibiotics. The plants that survived were planted in the soil to obtain sufficient material to check for transcript levels via PCR using primers specific to the TAP-tag. None of the plants tested amplified the genomic DNA of the insert. (Figure 5) At the present, large quantities of untested seeds are being grown along with newly acquired seedlings transformed using the same vector to compare germination rates at different concentrations of gentamycin.
Results of the Arabidopsis cell growth time course (Figure 6) revealed that cells enter exponential growth at four days after transfer. Using this data, it was decided to use cells at three days after transfer in order to optimize transformation.
Eight different transformations were attempted using cells grown in suspension culture cells. None produced lines that were transformed with the Tap-tag. Variations in the procedures tested were numerous. Gentamycin concentrations were lowered in the event that transformants were overly sensitive to gentamycin. Transfer conditions after the cocultivation were altered by reducing the number of washes and centrifugation in order to reduce the stress given to the cells. Growing media
was changed from Gamborg’s B5 to MS. Naphthalene acetic acid, a plant growth hormone, was added to the media to increase cell growth rate. Bacterial concentration was adjusted and decreased to prevent Agrobacteria from outgrowing the culture. One transformation tried an inoculation at 4 days after transfer instead of 3 days. Cells were grown in both the light and dark as well as shaking and stationary conditions. Finally, washed cells were returned to either fresh or conditioned liquid media or plated onto solid media. None of the sets of conditions produced viable cells after the exposure to gentamycin (i.e. no putative transformants were recovered from the gentamycin selection medium).
While waiting for transformed seedlings and cells to grow, a side project was initiated involving the characterization of a preexisting transgenic line of Arabidopsis (Hs9-7) that expressed a gene encoding the human form of the PIPK. The Hs9-7 is line known to have high levels of PtdIns(4,5)P
. (Im, et al. unpublished results).Permeability of control and transformed cells of WT and Hs9-7 line were tested by monitoring the uptake of FITC-Gln. Uptake was compared over time (from 1 to 6 h of incubation). Cells were photographed using bright field and fluorescence. Hs9-7 seedlings, previously described as having higher levels of PtdIns(4,5)P
, were consistently observed to have more fluorescence than WT plants.
WT roots screened had much lower fluorescence levels and the intracellular fluorescence was more diffuse in the WT seedlings. (Figure 7) These experiments show that altering the lipid composition might change membrane permeability or the activity of transporters.
Throughout the project, I’ve been working to optimize the transformation of Arabidopsis suspension cells with the TAP-tagged form of the PIP kinase. The techniques which I’ve learned throughout the project will make the production of transformants more likely in the near future. The newly acquired seeds from a neighboring lab which already contain the gentamycin resistance gene provide a positive control to optimize the gentamycin concentrations for selection. This will enable me to conduct the optimal gentamycin concentration to select but not kill the transformants, allowing for a higher throughput screening of seeds to begin with renewed focus and less uncertainty. In addition, new resources will be utilized to assist with the liquid cell culture project. I will be getting new protocol and advice from Richard Uhrig, a scientist who reported recent success at the American Society for Plant
Biologists meeting (August 2010). The potential key seems to be reducing the time between the transfer of the Arabidopsis cells and inoculation with the A. tumefaciens from 3 days to 2 days. This will increase the number of cells entering S-phase (DNA synthesis) of the cell cycle and should increase the probability for transformation. In addition, we will not wash the plant cells after co-cultivation to remove unbound A. tumefaciens before plating them on solid MS media containing timentin, an antibiotic that will kill the Agrobacteria. By increasing the efficiency at which the putative transformed seedlings can be screened and by using new techniques to transform cell cultures, I anticipate obtaining successful transformants in the near future.
Figure 1: Skematic model of the PI pathway. Decreased activity of the PIPK (highlighted in yellow), possibly because of regulation by interacting proteins, is predicted to be the cause of low basal levels of
in plant cells.
Figure 2: Model comparing the PIPK gene family across different types of species. The N-terminal MORN domain (labeled in green) is unique to plants and is predicted to be involved in regulation of the PIP kinase in subfamily B.
Figure 3A: Model of the TAP-tagged form of the PIPK. Attached to the kinase is an IgG binding domain, a
3C protease cleavage site, and a His binding domain which will be utilized to purify PIPK and PIPKinteracting proteins.
Figure 3B: Model of the dual-affinity purification method used to purify interacting proteins. The proteins of interest will then be further analyzed using mass spectroscopy.
Figure 4: Arabidopsis cell with bound A. tumefaciens (indicated with an arrow). Binding by the bacteria is required in order for the transfer of the Ti-plasmid into the plant cell and eventual incorporation into its genome.
Figure 5: Gel electrophoresis of PCR results. Samples run with primers specific to the TAP-tag (forward primer) and the IgG binding domain (reverse primer). A product of 515bp was predicted. The positive control, the recombinant plasmid DNA carries the TAP-tag construct, was the only lane to show amplification of the correct size (note arrow).
Day Day Day Day Day Day Day
0 1 2 3 4 5 6
Figure 6: Graph diagramming cell growth. Cells were transferred at two concentrations (1 and 2 mL of stock solution) and growth was measured daily. Cells were harvested by gravity filtration and fresh weight measured. Transformation should be optimal prior to exponential growth. We concluded that to obtain optimal cell transformation, cells should be used 3 days (arrow) after transferring 2mL of the stock lines.
Figure 7: Fluorescent images taken comparing uptake of FITC-glutamine between WT (left column) and
Hs9-7 (right column). Light-field images included to show comparison. Images taken after 14-day old seedlings were submerged in identical solutions of FITC-glutamine for four hours before being washed and imaged. The Hs9-7 line (bottom right) showed higher amounts of uptake than the WT line (bottom left), possibly due to higher levels of PtdIns(4,5)P
. Fluorescent images were exposed for the same time.
1. DeWald, D.B., et al. "Rapid accumulation of phosphatidylinositol 4,5-bisphosphate and inositol 1,4,5trisphosphate correlates with calcium mobilization in salt-stressed Arabidopsis." Plant Physiology 126
2. Forreiter, Christoph, Marc Kirschner, and Lutz Nover. "Stable Transformation of an Arabidopsis Cell
Suspension Culture with Firefly Luciferase Providing a Cellular System for Analysis of Chaperone Activity in Vivo." The Plant Cell 9 (1997): 2171-2181.
3. Im, Yang Ju, Amanda J. Davis, Imara Y. Perera, Eva Johannes, Nina S. Allen, and Boss Wendy F. "The Nterminal Membrane Occupation and Recognition Nexus Domain of Arabidopsis Phosphatidylinositol
Phosphate Kinase 1 Regulates Enzyme Activity." The Journal of Biological Chemistry, 2007: 5443-5452.
4. Im, Yang Ju, et al. "Increasing Plasma Membrane Phosphatidylinositol(4,5)bisphosphate Biosynthesis
Increases Phosphoinositide Metabolism in Nicotiana tabacum." The Plant Cell 19, no. 5 (May 2007):
5. Munnik, T., and Vermeer J. "Osmotic stress-induced phosphoinositide and inositol phosphate signalling in plants." Plant, Cell & Environment 33, no. 4 (April 2010): 655-669.
6. Perera, I.Y., A.J. Davis, D. Galanopoulou, Y.J. Im, and W.F. Boss. "Characterization and comparative analysis of Arabidopsis phosphatidylinositol phosphate 5-kinase 10 reveals differences in Arabidopsis and human phosphatidylinositol phosphate kinases." FEBS Letters 579 (2005): 3427-3432.
7. Perera, Imara Y., Chiu-Yueh Hung, Shari Brady, Gloria K. Muday, and Wendy F. Boss. "A Universal Role for Inositol 1,4,5-Trisphosphate-Mediated Signaling in Plant Gravitropism." Plant Physiology 140, no. 2
(February 2006): 746-760.
8. Rubio, Vicente, et al. "An alternative tandem affinity purification strategy applied to Arabidopsis protein complex isolation." The Plant Journal 41 (2005): 767-778.
9. Salinas, Julio, and Jose Sanchez-Serrano. Arabidopsis Protocols. Totowa, NJ: Humana Press, 2006.
10. Van Leene, Jelle, et al. "A Tandem Affinity Purification-based Technology Platform to Study the Cell
Cycle Interactome in Arabidopsis thaliana." ASBMB Molecular & Cellular Proteomics 6, no. 7 (April 2007):
11. Wiktorek-Smagur, A., K. Hnatuszko-Konka, and A. Kononowicz. "Flower bud dipping or vacuum infiltration—two methods of Arabidopsis thaliana transformation." Russian Journal of Plant Physiology
56, no. 4 (July 2009): 560-568.