Experiment 5: Transformation of Dictyostelium



One of the most important innovations in cell biology is the ability to genetically engineer cells to express genes of interest.  In this experiment, you will introduce the plasmid DNA into the cells by electroporation to transform the cells. 


Two plasmids will be used: one that expresses GFP (Green Fluorescent Protein) from a Dictyostelium promoter, and they one that expresses a fusion of GFP + the actin binding domain of Filamin. One will be a plasmid that integrates into the genome and one is a plasmid that replicates extrachromosomally.  Filamin is an actin crosslinking protein which possesses a 250 amino acid CH-domain that specifies its binding to actin filaments.  This domain has been engineered to be in frame with GFP so they are expressed as a single fusion protein.  In cells, this GFP fusion probe localizes with the actin cytoskeleton.


Questions:

1. What is the efficiency of transformants for each vector?

2.How does the extrachromosomal vector differ from the integrating vector in terms of proportion of cells that express GFP.

3. How does the localization of GFP compare to the localization of GFP-FilABD.  What do you expect to see for each? 


You will need lots of cells for this experiment.  Split your growth plates accordingly so you have about 1plate per cuvette you are going to do


Procedure:


Setting up the experiment


Each group will prepare 3 sterile cuvettes:     


1)no DNA control

2)PDM-GFP extrachromosomal vector

3)GFP-Fil-ABD vector


Label your cuvettes and put them on ice to chill. 

Set up the Petri dishes to put the cells in after electroporation (step 8)


Calculate what volume of DNA you will need for each cuvette. You will need 4 µg of DNA per cuvette.


The procedure does not require a specific number of cells or amount of DNA.  Generally, the more cells and DNA you use, the more transformants you will get up to some (unknown) limit.



Harvesting AX2 Cells in H50 Buffer


1.  We are using Ax2 for this experiment because you can actually count colonies of transformants.  NC4A2 is so motile, that colonies never form, so it is much harder to assess the transformation results. 


2.  Harvest your Ax2 cells into a 15 ml centrifuge tube on ice. Take an aliquot to titer on the hemocytometer (so you know how many cells you are spinning down).  Centrifuge in 15ml conical tube for 5 min (1000 rpm) at 4°C.


THE REST OF THE ASSAY IS DONE ON ICE! Cells and buffers must be kept cold throughout this procedure  IT IS IMPORTANT TO KEEP THE CELLS COLD AND TO NOT TO LEAVE CELLS IN H50 BUFFER FOR ANY LONGER THAN NECESSARY – THE HIGH SALT WILL KILL THE CELLS!


3.After spinning, carefully aspirate or pour off the supernatant leaving the pellet untouched and resuspend the cells in 5ml ice-cold H50 buffer. It works best if you finger flick the tubes (ask how to do this if you don’t know) to dislodge the pellet before adding the buffer.


4.Spin the cells again for 5 min (1000rpm) at 4°C.


5.While cells are spinning, calculate to volume of H50 buffer to resuspend your cells in to give a titer of 1-5 x 107 cells/ml.


6. Aspirate H50 from the tubes and add that amount of cold H50 to your cells and resuspend the pellet.


7.In a microcentrifuge tube on ice, mix 4µg DNA, cells and bring the total to 400 µl H50 buffer  and gently mix


8.Transfer the mixture to the ice cold 0.2 cm cuvettes. Do this shortly before you have access to the electorporator.  If not, the cells will have settled to the bottom of the cuvette before you get to pulse them. 



Electroporation


1. Bring entire ice bucket to the electroporator. We will be doing electroporation using 2 different settings. Half the class will do one and half the class the other.


a.Exponential setting: 700 V, 50 Ω, 50 uF, 0.2 mm cuvette; 2 pulses 1 minute apart.


b.Time constant setting: 700 V, 0.6 ms, 0.2 mm cuvette; 2 pulses, 1 minute apart.



The TA will assist you to choose the program to use in the Home menu of the electroporator.   The two conditions are programmed into memory.  Pulse the cells, then put the cuvette back on ice and pulse the other cuvettes.  Then repeat the pulsing.  There should be about 1 minute between pulses, but the time is not critical.  Return the cuvettes to the ice bucket and incubate on ice for 5 minutes. Take note of the time constant for each pulse and write this data in your notebook.



2. Transfer the cells from each of the 3 cuvettes (with a pipetman) to their corresponding labeled Petri dish with 10ml of HL5. By tilting the cuvette and sticking your yellow tip down the side, you can remove most of the volume in the cuvette.  To remove the rest of the cells, take 100 µl of HL-5 from that same Petri dish and add it to the cuvette, pipette up and down a couple of times, and then remove it from the cuvette and add it back to the plate with the cells.  Now disperse the cells in the dish by swirling.  Remove 1 ml of the cells plus media and add it to a second dish with 9 ml of HL5.  Add 1ml of HL5 to the original plate to bring it to 10 ml. This will give you two dishes to look at for transformation efficiency.  If there are too many colonies to count on the high titer dish, then the diluted plate should give you 1/10 as many colonies.


3.Washing and sterilizing the cuvettes for future use.  They can be used many times if treated appropriately.

a.When you are done with the cuvettes, rinse them about 5-10 times with water from the tap or squirt bottle, flicking out the water in between.

b. Fill the cuvette with 70% ethanol from the squeeze bottle

c. Put the cap on and invert several times

d. Pour off the ethanol and put the cap back on. 

e. Place the clean sterile cuvette back in the rack to dry.



For your Notebook/web site, record an image of what one of the plates looks like at this point.  Presumably, they will all look the same, but if not, note which looks different and why.  Did the electroporation leave them rounded, dead looking or do they look healthy? Are the numbers of cells the same for the different conditions (ie, did one procedure seem to kill more cells than the other?)



The Next Day


1.  The next day (precise time is not critical, but record it), add 5ul 1000x G418 (10mg/ml) to each plate (final concentration 5 µg/ml).  This will kill any cells that did not integrate the antibiotic resistant DNA plasmid into their genome.


2.  Over the next few days you will see sensitive cells first round up and then detach from the plate as they die.  Resistant cells will remain attached and grow as colonies.  By day 3, they should be mostly detached.  Take an image each day to record what is happening in the dishes. You should not need to take an image of each plate presuming they all look basically the same.


Day 3


1. On about day 3, most of the cells will have rounded up and detached from the surface(they are dead), remove the dead cells by gently swirling the plate to get the dead cells in suspension.  The dead cells form a layer on the bottom of the dish and you can see them become suspended as you swirl.  Check in your benchtop microscope to see you have done a good job and then aspirate the media and dead cells.  Add back 10 ml fresh HL5 media and 5 µl G418.  Check the plate to see how well you did in getting the dead cells out.  If there are still too many, you could repeat the wash but this is not usually necessary. 


2. Carefully check the plate for colonies of transformed cells growing on the dish (you can check before as well by swirling and then looking in the microscope).  If you find them, take some images.  You can also look using the blue LED illuminator to look for fluorescent cells.  Let the cells sit in fresh media and drug for 5-7 days and keep monitoring the colonies. 


IMPORTANT: BE GENTLE IN SWIRLING. YOU don’t want to detach your TRANSFORMANTS from the bottom of the dish!!!  You just want to get the dead cells off the bottom.


3. After 5-7 days or longer, LOOK at your cells under fluorescence microscope scope to see fluorescent transformants.  Some cells will be expressing GFP. What proportion of the cells express GFP?  Remember that all of the cells must possess the plasmid in order to grow in the presence of the antibiotic.  As the colonies grow, you can hold the plate up to the light and see them by eye.  Each colony represents a single cell that retained the plasmid and grew into a colony of cells.  Where does the probe localize?  Can you count the colonies to assess transformation efficiency?  What proportion of the cells are fluorescent?


4. Once you have these cells, keep them growing as you may need them for future experiments.



Lab Writeup:


Keep notes on everything (including calculations, observations and expectations) in your notebook and post images of your transformants when available in your website with a brief description. In your writeup, discuss what happened during the various phases of the experiment and your results.  Give the transformation efficiencies of each plasmid and electroporation condition and the frequency of GFP positive cells.  Did the control work? 



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Reagents/Media:

HL5 media

H50 Buffer

G418 Drug


Cell Strain(s):

Ax2


Other things needed for this experiment:

1.Electroporator

2.sterile 0.2 cm cuvettes

3.Ice buckets and lots of ice

4.15-ml blue-capped falcon tubes

5.colored tapes for labeling cuvettes

6.Centrifuge should be on and set at 4oC


Add electroporation buffer recipe

G418 stock recipe

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