SBB09Ntbk-Roger Lowe

From OpenWetWare

Revision as of 15:46, 6 May 2009 by W. Roger Lowe (Talk | contribs)
(diff) ←Older revision | Current revision (diff) | Newer revision→ (diff)
Jump to: navigation, search

Contents

Silver Binding Assay

Wednesday: April 29, 2009

Results from incubation on Monday:
All of the tubes had some sort of reddish precipitate except the control. Construct 13 had the most precipitate and 16 had the least. 11 and 16 which did not have precipitate in the first experiment had precipitate.
tried to resuspend the reddish particles, but did not resuspend despite lots of vortexing and other attempts.
tried to - 1. add lysozyme 2. manually remove precipitate and put into water 3. lots of vortexing
Took absorbance measurements of control, 16 (lysozyme), 15 (in water), and 13 (vortexing). After normalizing the graph, the control had least absorbance, but there is no significant difference in signal in the 400-440nm range, so inconclusive results.

grew cells in 4ml LB and 4ul arabinose. used DH10B as control instead of 1363.

method to resuspend reddish particles (assumed to be silver nanoparticles): 
1. pellet cells and particles
2. pour out excess TBS and AgNO3
3. wash with water 2X and resuspend
4. react Ag (red particles) with nitric acid
5. react Ag ions with KBr. KBr(aq) + AgNO3(aq) → AgBr(s) + KNO3(aq)
6. soluble yellow product should form
7. measure absorbance of the yellow product
8. quantify the amount of silver (of the different constructs) formed from the intensity of the color.


Monday:April 27, 2009

resuspended cells and incubated them in TBS and 0.1mM AgNO3 (new) at room temperature. put on rocker to agitate the solutions. (repeated experiment)

measured absorption of the various samples using the Tecan.
resuspended the pellets, but chunks of cells remained in solution. pipeted 100ul of each sample into column 9 of plate.
saved data as spectro ver1, ver2, etc
parameters: 250-800nm range. 10nm intervals. 96 well plate.

{graph of absorbance vs wavelength}

graph did not indicate differences between the control and samples with color. There could be too much scattering. could solubilize with base or proteinase to make solution clear. Alternatively, could get rid of cells and measure the total amount of silver bound to cells after washing out excess silver.


Sunday: April 25, 2009

checked the cells. noticed a noticeable color change (a reddish color) in the pellet for some of the tubes.

control 1363 - white
11 - white {Pbad.rbs.prepro.StrepTag}{<AG4>}{<CPG_L6!}{dblTerm}
12 - reddish* {Pbad.rbs.prepro.StrepTag}{<AG4>}{<eCPX!}{dblTerm}
13 - reddish {Pbad.rbs.prepro.StrepTag}{<AG4>}{<upaG_short!}{dblTerm}
14 - reddish {Pbad.rbs.prepro.StrepTag}{<AG4>}{<Ag43_short!}{dblTerm}
15 - reddish {Pbad.rbs.prepro.StrepTag}{<AG4>}{<espP(beta)!}{dblTerm}
16 - white {Pbad.rbs.prepro.StrepTag}{<AG4>}{<ehaB!]{dblTerm}
17 - reddish {Pbad.rbs.prepro.StrepTag}{<AG4>}{<CPompX!}{dblTerm}

*deepest color change

grew more cells in the presence of arabinose (4ml LB, 4ul of 1000X arabinose, constructs 11-17 with control)

 
need to: 
1. repeat the experiment
2. find a way to quantify the color change - look in literature 
3. take picture of the solutions

Monday:
will do -
repeat the same experiment done on Friday
search literature for way to quantify results
carry out experiment in 96 well plate with more variations

So far:
1. when grown in presence of silver nitrate, no cell growth
2. when grown in the absence of arabinose, washed twice with TBS, and resuspended in 0.1mM AgNO3 and TBS, no color change in washed pellets
3. when grown in presence of arabinose, washed twice with TBS, and resuspended in 0.1mM AgNO3 and TBS, color change observed in washed pellets

Friday: April 24, 2009

washed (2X) the cells grown on Thursday in 200ul of TBS. Both the TBS and 1mM AgNO3 solutions were freshly made.
resuspended each of the pellets in 180ul of TBS and 20ul of AgNO3 (to make 0.1mM AgNO3)




Thursday: April 23, 2009

The cells grown in AgNO3 on Tuesday seemed to have died. Perhaps the pH change killed the bacteria.

We washed the cells (we grew on Tues) with TBS, put them in 180ul of TBS and 20ul of 0.1mM AgNO3, and then put them on the shaker. no color change and not much apparent change.

We also grew the washed cells in 3ml LB and 0.1mM AgNO3
No cells seemed to have grown. silver is toxic to cells.

Chris put in formaldehyde to the cells. Cells should have turned black but didn't. not sure if enough silver nitrate was added.

Regrew cells in the presence of arabinose. Constructs 11-17 + control.
So hopefully, the cells will express the silver binding peptides and then on Friday, the TBS and the silver nitrate will be added.


Wednesday:April 22, 2009

 make TBS, make AgNO3, measure OD 

cells of var. conc., arabinose, AgNO3 of var. conc., TBS


growing cells grid

Tuesday: April 21, 2009

grew one colony from each sample (11-17) and control in 4ml LB (with CA)
grew one colony from each sample (11-17) and control in 4ml LB (with CA) and 0.1mM AgNO3

Monday: April 20, 2009

transformed constructs into bacteria
plated the cells
we will use constructs 11-17 and a pBca9145-Bca1363 control




The cells grown in AgNO3 on Tuesday seemed to have died. Perhaps the pH change killed the bacteria.

We washed the cells (we grew on Tues) with TBS, put them in 180ul of TBS and 20ul of 0.1mM AgNO3, and then put them on the shaker. We also grew the washed cells in 3ml LB and 0.1mM AgNO3

EeaA Intimin gene part construction

February 18, 2009
Created cloning solution for EaeA Intimin gene using buffer, primers, expand polymerase, and template DNA.

PCR reaction

*Make 100uM stocks of primers, this is the concentration used directly for the reaction
*Prepare the following reaction:
  29 uL water
  5 uL Expand 10x Buffer 2
  5 uL 10x dNTPs (2 mM in each; 0.2 mM final conc)
  5 uL Oligo 1 (100uM)
  5 uL Oligo 2 (100uM)
  0.75 uL Expand Polymerase 1


Ran on gel to check size/ presence of EaeA gene. The gene ran slightly further than expected, suggesting that it could be smaller than the hoped for gene.

February 25, 2009

Regular Zymo Cleanup

The following procedure removes the polymerase, dNTPs, buffer, and most of the oligonucleotides from a PCR reaction. It also will remove the buffer and restriction enzymes from a restriction digest reaction.

#Add 180 uL of Zymo ADB buffer (brown bottle) to a 33uL or 50uL reaction.
#Transfer into the Zymo column (small clear guys)
#spin through, discard waste.
#Add 200 uL of PE or Zymo Wash buffer (which is basically 70% ethanol)
#spin through, discard waste.
#Add 200 uL of PE or Zymo Wash buffer
#spin through, discard waste.
#spin for 90 seconds, full speed to dry.
#elute with water into a fresh Eppendorf tube, use the same volume of water as the volume of the original reaction

During this purification I put my reaction tube in the centrifuge unlabelled and it got confused with Nihkeil. We reran our samples side by side on a gel, and found which sample was my EaeA part.

Electrophoresis

2ul EaeA PCR sample
1ul Dye


February 27, 2009
In our gel results, my protein ran to about 1.7 kb, Nikhil's did not show, so we concluded that since my gel ran to about the expected length of my gene, I had grabbed my gene, EaeA intimin. I continued on to digesting and ligating my gene to the pBca9495CA plasmid.

EcoRI/BamHI Digest of PCR Products

For PCR products, you will only digest a portion of your purified PCR product. Note that you must make a minor modification to the procedure if your DNA is shorter than 300bp (you add a little isopropanol to the ADB after melting).

  • Set up the following reaction:
 8uL of eluted PCR product
 1uL of NEB Buffer 2
 0.5uL EcoRI
 0.5uL BamHI
  • Incubate at 37 degrees on the thermocycler for 1hr
  • Run an agarose gel, and melt with 600uL ADB buffer at 55 degrees. ****NOTE: If you are running short of time, this is an acceptable stopping point
  • If the DNA is shorter than 300bp, add 250uL of isopropanol and mix prior to loading it on the column

Ligation of EcoRI/BamHI digests

*Set up the following reaction:
  6.5uL ddH2O
  1uL T4 DNA Ligase Buffer (small red or black-striped tubes)
  1uL Vector digest
  1uL Insert digest
  0.5uL T4 DNA Ligase
*Pound upside down on the bench to mix
*Give it a quick spin to send it back to the bottom of the tube
*Incubate on the benchtop for 30min
*Put on ice and proceed to the transformation

Transformation by heat-shock

Competent cells are stored as 200uL aliquots in the -80 freezer as a communal stock.

#Thaw a 200 uL aliquot of cells on ice
#Add 50 uL of water
#Add 30 uL of KCM salts
#Put your ligation mixture on ice, let it cool a minute or two
#Add 75 uL of the cell cocktail to the ligation, pipette up and down gently to mix
#Let sit on ice for 10 min
#Heat shock for 2 min at 42
#Put back on ice for 1 min
#For ampicillin selection, you can plate immediately, otherwise:
#Add 100uL of LB, let shake in the 37 degree incubator for 40 min
#Plate on selective antibiotics, let incubate overnight

Picking of colonies

    * For each construct you will pick and later miniprep 2 colonies
    * Add 4mL of LB media with the appropriate antibiotics to a clean test tube
    * Pick a well-isolated, round, and "normal" looking colony with a toothpick
    * Drop it in the test tube
    * Incubate at 37 overnight 

Miniprep purification of DNA

MINIPREP (1mL - 5mL) Procedure for Plasmid DNA Purification (using the QIAGEN QIAPrep Spin Miniprep kit)

#Pellet around 1.5 mL or 2 mL saturated culture by spinning full speed, 30 seconds.
#Dump supernatant, repeat to pellet another 1.5 mL (for a total of 3 mL)
#Add 250uL of P1 buffer into each tube. Resuspend the cells using a vortexer.
#Add 250uL of P2 buffer (a base that denatures everything and causes cells to lyse). Gently mix up and down. Solution should become clearer.
#Add 350uL of N3 buffer (an acid of pH ~5 that causes cell junk - including protein and chromosomal DNA - to precipitate, and leaves plasmids and other small molecules in solution). Slowly invert a few times, then shake.
#Spin in centrifuge at top speed for 5 minutes.
#Label blue columns with an alcohol-resistant lab pen.
#Pour liquid into columns, and place the columns into the centrifuge. Spin at 12000 rpm for 30 seconds.
#Dump liquid out of the collectors under the columns (the DNA should be stuck to the white resin)
#Wash each column with 500 uL of PB buffer.
#Spin in centrifuge at 12000rpm for approximately 15 seconds, then flick out the liquid again.
#Wash with 750uL of PE buffer (washes the salts off the resins).
#Spin in centrifuge at 12000rpm for approximately 15 seconds and flick out liquid again.
#Spin in centrifuge at full speed for 1 minute to dry off all water and ethanol.
#Label new tubes and put columns in them.
#Elute them by squirting 50uL of water down the middle of the column (don't let it stick to the sides).
#Spin in centrifuge at top speed for 30 seconds.
#Take out columns and cap the tubes.
#Clean up - note the P1 buffer is stored at 4degC and all the rest at room temperature.


March 2, 2009
I did not get any colonies for EaeA-CA transformed plasmid. I started again at ligation with the digested and cleaned EaeA plasmid. My ligation and transformation reactions followed the same procedures as the ligation and transformation protocol listed above.
I also started a new EaeA intimin PCR reaction in case my second plating efforts do not produce any colonies, using the same PCR protocol as explained above.


My second set of transformations for EaeA plasmid produced colonies. I performed miniprep with the same protocol as above. I performed plasmid ligation for mapping and then ran on a gel.


March 6, 2009
I sequenced my piece and it turned out that Nikhil and I had actually switched are reactions in the centrifuge. The sequence I got was an exact match for Nikhil's TraT part. I checked to see how his sequence matched with mine and it was a bad read. I got his second colony miniprep sequenced and it showed a fragment of my EaeA gene. We conjecture that the primers overlapped in this circumstance producing only part of the gene. I should have gel purified to get the portion of the reaction that had my desired gene size.

Aluminum Binding Protein gene part construction

February 18, 2009
I started my wobble reaction with the oligos designed and ordered the week before.

Wobble Reaction

The Wobble procedure is a variation of Klenow Extension that begins with two oligonucleotides that overlap by around 20 bp on their 3' ends and uses a thermostable polymerase

  • Order the oligos, they don't need to be purified in any special way, smallest scale is ok
  • Make 100uM stocks, this is the concentration used directly for the reaction
  • Prepare the following reaction:
  
29 uL water
  5 uL Expand 10x Buffer 2
  5 uL 10x dNTPs (2 mM in each; 0.2 mM final conc)
  5 uL Oligo 1 (100uM)
  5 uL Oligo 2 (100uM)
  0.75 uL Expand Polymerase 1
  • Run the wobble program, whick is:
 2 min at 94
 10 cycles of:
 30 sec at 55
 30 sec at 72
 (or something similar)
  • There is no point in running an analytical gel afterwards, there is nothing to see
  • You'll want to run short fragment cleanups to remove the polymerase prior to digestion steps


February 23, 2009
After performing Klenow extension for aluminum binding gene, I performed Zymo Cleanup.

Small-Frag Zymo Cleanup

The following procedure removes the polymerase, dNTPs, buffer, and most of the oligonucleotides from a PCR reaction for fragments smaller than 300bp. It also will remove the buffer and restriction enzymes from a restriction digest reaction.

#Add 100 uL of Zymo ADB buffer (brown bottle) to the reaction.
#Transfer into the Zymo column (small clear guys)
#Add 500uL of Ethanol and pipette up and down to mix
#spin through, discard waste.
#Add 200 uL of PE or Zymo Wash buffer (which is basically 70% ethanol)
#spin through, discard waste.
#Add 200 uL of PE or Zymo Wash buffer
#spin through, discard waste.
#spin for 90 seconds, full speed to dry.
#elute with water into a fresh Eppendorf tube

Now to be cyclized

Created solution for Klenow (wobble) extension for aluminum binding protein, now being cyclized.
I then continued to digest the aluminum binding peptide gene.

EcoRI/BamHI Digest of Wobble Products

For wobble products, you will digest the entire extension reaction-worth of DNA Since you started with 50uL of extension reaction, you should now have 50uL of eluted DNA in water.

 50uL eluted DNA
 5.7uL NEB Buffer 2
 1uL EcoRI
 1uL BamHI
   * Mix thoroughly by slamming the tube upside down on the table, and then a quick spin to move the liquid to the bottom of the tube.
   * Incubate the reaction at 37 degrees on the thermocycler
   * Proceed to another Zymo small fragment cleanup

February 25, 2009

Ligation of EcoRI/BamHI digests

  • Set up the following reaction:
  
6.5uL ddH2O
  1uL T4 DNA Ligase Buffer (small red or black-striped tubes)
  1uL Vector digest
  1uL Insert digest
  0.5uL T4 DNA Ligase
  • Pound upside down on the bench to mix
  • Give it a quick spin to send it back to the bottom of the tube
  • Incubate on the benchtop for 30min
  • Put on ice and proceed to the transformation

Transformation by heat-shock

Competent cells are stored as 200uL aliquots in the -80 freezer as a communal stock.

#Thaw a 200 uL aliquot of cells on ice
#Add 50 uL of water
#Add 30 uL of KCM salts
#Put your ligation mixture on ice, let it cool a minute or two
#Add 75 uL of the cell cocktail to the ligation, pipette up and down gently to mix
#Let sit on ice for 10 min
#Heat shock for 2 min at 42
#Put back on ice for 1 min
#For ampicillin selection, you can plate immediately, otherwise:
#Add 100uL of LB, let shake in the 37 degree incubator for 40 min
#Plate on selective antibiotics, let incubate overnight

Picking of colonies

  • For each construct you will pick and later miniprep 2 colonies
  • Add 4mL of LB media with the appropriate antibiotics to a clean test tube
  • Pick a well-isolated, round, and "normal" looking colony with a toothpick
  • Drop it in the test tube
  • Incubate at 37 overnight


February 27, 2009
My two colonies for ABP in 9495-CA plasmid were labeled bomb 1 and bomb 2. I mini-prepped both of them.

Miniprep purification of DNA

MINIPREP (1mL - 5mL) Procedure for Plasmid DNA Purification (using the QIAGEN QIAPrep Spin Miniprep kit)

#Pellet around 1.5 mL or 2 mL saturated culture by spinning full speed, 30 seconds.
#Dump supernatant, repeat to pellet another 1.5 mL (for a total of 3 mL)
#Add 250uL of P1 buffer into each tube. Resuspend the cells using a vortexer.
#Add 250uL of P2 buffer (a base that denatures everything and causes cells to lyse). Gently mix up and down. Solution should become clearer.
#Add 350uL of N3 buffer (an acid of pH ~5 that causes cell junk - including protein and chromosomal DNA - to precipitate, and leaves plasmids and other small molecules in solution). Slowly invert a few times, then shake.
#Spin in centrifuge at top speed for 5 minutes.
#Label blue columns with an alcohol-resistant lab pen.
#Pour liquid into columns, and place the columns into the centrifuge. Spin at 12000 rpm for 30 seconds.
#Dump liquid out of the collectors under the columns (the DNA should be stuck to the white resin)
#Wash each column with 500 uL of PB buffer.
#Spin in centrifuge at 12000rpm for approximately 15 seconds, then flick out the liquid again.
#Wash with 750uL of PE buffer (washes the salts off the resins).
#Spin in centrifuge at 12000rpm for approximately 15 seconds and flick out liquid again.
#Spin in centrifuge at full speed for 1 minute to dry off all water and ethanol.
#Label new tubes and put columns in them.
#Elute them by squirting 50uL of water down the middle of the column (don't let it stick to the sides).
#Spin in centrifuge at top speed for 30 seconds.
#Take out columns and cap the tubes.
#Clean up - note the P1 buffer is stored at 4degC and all the rest at room temperature.


March 02, 2009
I digested my isolated plasmids using EcoR1 and BamH1.

Plasmid Digestion

5ul dH20
3ul DNA solution
1ul NEB buffer
0.5 ul EcoR1
0.5 ul BamH1


Both of my samples showed plasmid backbones that ran on the gel, which indicates that the plasmids did indeed transform into the cells.

March 06, 2009
I entered my ABP plasmid information in the sequencing and stock log, and inserted 20ul into the sequencing plates at H2 for plate 1 and 2.

March 09, 2009
My sequencing data was a perfect read for my part.

Personal tools