Protocols for Yeast CRISPR CURE

This is unpublished work in progress and if you’re using, we’re happy to incorporate any of your feedback.

Email bwasko@westernu.edu

Brief introduction to the lab.

About course-based undergraduate research experiences (CUREs):This lab is designed to be a course-based undergraduate research experience.  That means the student will be developing a novel hypothesis and performing authentic scientific research with an unknown outcome.  Using CRISPR, students will genetically engineer the genome of Saccharomyces cerevisiae (Baker’s yeast) to test their own hypothesis related to the structure and function of an alkaline phosphatase enzyme (Pho13).  A human homolog is glycerol-3-phosphate phosphatase (PGP).

Protocols for each technique are found below as ‘Labs’. This was initially developed for a biochemistry laboratory, but in theory you could pick any gene(s) and follow labs 0-6 which could feed into many genes/phenotypes/assays of your choosing.

Prior to starting experiments, oligos will need to be designed and ordered.  Allow for time for hypothesis generation, and oligo design and delivery.  Working in large groups for hypothesis generation can reduce reagent requirements, TA/instructor experimental logistics, and redundancy in the event of individual student experiment failure. Some labs were initially structured to be 2 hour labs, but some labs could easily be combined to reduce the number of lab days necessary to complete the CURE if more lab time is available.

See also UHCL Biochem Lab CRISPR CURE version used by Dr. Scott Buckel (forked when BW left UHCL)

CRISPR background

CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats) was discovered as part of an antiviral immune system that some bacterial contain.  After exposure to a virus, these bacteria can incorporate segments of the viral genome into their own DNA, which can allow for a memory of the virus to be used in the event of re-exposure to the virus. This DNA is made into RNA that works with a bacterial CRISPR associated protein, such as CAS9.  The RNA helps to target the CAS9 protein to the complementary viral DNA so that the CAS9 endonuclease enzyme can cleave the DNA. CAS9 cuts DNA on both strands, leading to double strand breaks (DSBs).  Scientists, including those in the laboratories of Dr. Jennifer Doudna and Dr. Emmanualle Charpentier, who received a Nobel Prize in 2020, engineered two RNA sequences from this bacterial system into a single sequence (single guide RNA or sgRNA).  The sgRNA contains ~20 nucleotides that are complementary to the target DNA that is to be cut.  The CAS9 enzyme and an sgRNA are all that are required for creating a highly sequence specific double strand break in the genome.  This system can be employed to cost effectively, easily, and specifically edit a cell's genome.  

The CRISPR/CAS9 system (Figure 1) requires a protospacer adjacent motif (PAM) sequence present in the targeted DNA in order to cut the DNA 3 bp upstream of the PAM site. This PAM sequence for Cas9 is NGG (where N is any nucleotide) and follows the complementary targeting of the sgRNA.  The DNA encoding for the sgRNA lacks the NGG sequence, which allows for the CRISPR/CAS9 system to not cut the DNA that encodes itself, but instead to cut only where the PAM sequence is present.

Figure 1. Cas9 and sgRNA. Image from Wikimedia commons. Mariuswalter, CC BY-SA 4.0

Cells will arrest or die if they are unable to repair a DNA double stranded break. There are two pathways cells use to repair DNA double-stranded breaks.  Homology directed repair (HDR) uses a homologous sequence as a template to repair the broken DNA with high fidelity.  Non-homologous end joining (NHEJ) does not use a homologous sequence for repair and is more error prone.

CRISPR yeast CURE background

In order to deploy CRISPR/CAS9 in yeast, you will use a plasmid already made by a research lab (Laughrey et al. Yeast. 2015) that encodes for the Cas9 protein and has the sgRNA only lacking the targeting sequence.  The targeting sequence can be cloned into the plasmid using a classical restriction enzyme cloning technique.  Since this is a small sequence, custom synthesized oligonucleotides (oligos) can be cost-effectively purchased from a company and then used for the cloning.

You will make one set of oligonucleotides that can be used to clone the sgRNA targeting sequence into the plasmid pML104.  Restriction enzyme sites for cloning are included in the sequence when generated from the authors’ website informatic tool and are also posted on blackboard.  Then you will make a second set of oligos (that we will refer to as the 'repair template') which will consist of two complementary oligonucleotides ~60bp long.  This sequence will be homologous to the sequence surrounding the targeted PAM site and desired mutation site that you will be making.  The repair template will contain a silent PAM mutation and a desired mutation (Fig.2).

Following the cloning, the resulting plasmid will have CAS9, the sgRNA with targeting sequence cloned, and the URA3 (uracil biosynthesis) gene.  The laboratory yeast strain used lacks the URA3 gene, so growth in the absence of uracil in the growth media is a positive selection for only yeast that transformed (took up) the plasmid.  However, having Cas9 with the sgRNA is then a negative selection so yeast that transform with the plasmid will not survive due to double strand breaks at the targeted site in the genome.  Cells will not progress through the cell cycle with double strand breaks in the genome. If cells repair the damage perfectly, CAS9 w/the sgRNA would simply recut the DNA again.  If yeast use the 'repair template' oligos as a template for repair of the double strand breaks via homologous recombination, then they will incorporate the silent mutation to the PAM site so CAS9 will no longer cut DNA and these cells will survive, and then the mutation of interest will have also been inserted.

Experimentally, you can get an indication that the genome modification experiment worked, based on colony counts alone.  Yeast transformed with plasmid containing CAS9 + your sgRNA alone should yield zero or very few colonies, and when the repair template is included ideally it should increase to significantly more colonies.  The parental plasmid with no sgRNA targeting sequence can be used as a positive transformation control to verify the yeast transformation is working well (>100s of colonies).  Yeast with no plasmid transformed can be included as a negative transformation control (zero colonies).  The gene will then be PCR amplified and sequenced to determine if the desired mutation is present.  Then the mutant strain will be grown, the enzyme will be isolated, and your hypothesis will be tested by assaying enzymatic activity.

Figure 2. Cloning and CRISPR/Cas9 strategy in yeast. gDNA = genomic DNA and ORF = Open Reading Frame of gene.  

PHO13 background

        The yeast PHO13 gene encodes for the Pho13 protein, which is a conserved alkaline phosphatase enzyme.  It has an alkaline pH optimum and has dephosphorylation activity, including the ability to dephosphorylate 2-phosphoglycolate.  Pho13 has homology to human  glycerol-3-phosphate phosphatase (PGP).  The enzymatic activity of Pho13 can be measured using the colorless substrate para-nitrophenyl phosphate (PNPP) which is converted by Pho13 to the yellow (absorbance at 400nM) compound para-nitrophenol.

BIOL4242 students see Assignment 1 on blackboard and Sequence alignment video

BIOL4242 students see Assignment 2 Hypothesis generation and oligo design tutorial

Lab 0. Oligo Design

*Be sure to check out the oligo design video 

Note that you will use PHO13, not TRP1.

• crisprhelper.wasko.org

• The entire process below is automated in the above software link.

Determine the codon to alter to create your mutation of interest

a)      Find where your desired mutation is in the amino acid sequence and DNA sequence of your gene/protein (e.g., Pho13).

b)      Download the yeast DNA coding sequence from SGD yeastgenome.org

c)       Open the file in Ape (A plasmid editor)

d)      Translate the sequence in ApE

a.       highlight everything in the sequence (if you have just the full open reading frame of the gene),  or select ‘ORF’ – ‘Find next’

e)      ‘ORF’ ‘Translate’

a.       select ‘codon spacing’, ‘line numbers’ ‘both’, ‘DNA above’

f)        Determine the DNA sequence that corresponds to the amino acid residue you want to mutate.

a.       Consider changing the case (‘Edit’ – ‘Upper <-> Lower’ or on PC press Ctrl +) or use highlighting by selecting ‘Features’ ‘new feature’.

g)       Using a codon table, change as few nucleotides as possible to alter the codon to one that encodes for your desired amino acid residue(s).  Change the DNA sequence to include your desired mutation.

Design oligonucleotides necessary to clone into a vector that would target the genome near where you desire your mutation

a)       For background, consider reading the Wyrick lab published article for additional information

b)      First find a gRNA target site in the yeast gene of interest as near to your desired mutation site as possible.  

Full list of Pho13 target sites from Wyrick lab output.  Uppercase indicates sgRNA targeting region. These ‘oligo1’ and ‘oligo2’ sequences will already contain the cloning sites mentioned above e.g., GATCNNNNNNNNNNNNNNNNNNNNGTTTTAGAGCTAG

Note that some sequences can show the complementary sequence in the genome (CCNNNNNN, where CC is complementary to the GG of the PAM site). At the end of the gRNA targeting sequence (GG could be CC as other strand) will be the PAM site.  Note that the PAM site NGG is NOT located in the cloning oligos, otherwise CAS9 would cut the cloned gRNA coding DNA in the plasmid. If you search for your gRNA targeting sequence you should find it (or the reverse complement) in your cloning oligos (without the PAM site).

Alternatively, to find sgRNA targeting sites in the yeast genome,

• crisprhelper.wasko.org

Backup python script Yeast AutoOligo CRISPR Helper 1.5 Colab (google.com)

        Be sure your sequences contain append the following sequence with your sgRNA targeting region to make your oligos:

        Oligo 1:GATCNNNNNNNNNNNNNNNNNNNNGTTTTAGAGCTAG

        N=CRISPR site (without PAM)

        Oligo 2: CTAGCTCTAAAANNNNNNNNNNNNNNNNNNNN

        N=CRISPR site reverse complement (without PAM)

.

c)       Copy and paste the gRNA target sequence, and Oligo1 and Oligo2 sequences into a word processor document.  These already contain necessary restriction enzyme sites to clone them into a CRISPR/CAS9 containing plasmid that can be transformed into yeast cells.

d)      Determine where the NGG of the PAM site is in regards to the open reading frame, and using a codon table, determine if you are able to make a nucleotide substitution(s) to change at least one of the GG nucleotides (or CC if gRNA sequence is on other strand) to a different nucleotide(s) to make a silent mutation (a different codon that still encodes for the same amino acid residue).  Record the nucleotide change you are making.You may run into the case (e.g., if GGN is codon) where you are unable to change a G in the PAM site to make a silent mutation.  Just restart and choose a different guide RNA sequence if that occurs.

e)      From the gene (PHO13) DNA sequence, select 60 bp of sequence that surrounds your PAM site and mutation site (trying to center in the sequence the PAM mutation and your mutation as best you can).  Include the PAM silent mutation and the desired mutation in this 60bp sequence.  Store this sequence and name it RepairTemplate.  With this same sequence, select ‘Edit’ then ‘Reverse Complement’ within ApE to get the complementary strand of DNA to this sequence, and call this the other strand of DNA for the repair template. These two oligos can be annealed to form double stranded DNA.

order oligos from supplier such as ThermoFisher Scientific or alternative

Lab 1 Cloning guide sequence of sgRNA

Background video

Molecular cloning is the process of piecing together pieces of DNA from multiple sources.

Equipment and Materials (per group):

·         TOP10 or similar E.coli (3 tubes/student) (homemade competent, make and test transform first)

·         LB+100µg/ml Ampicillin plates (3-6 plates) (or Carb instead of Amp)

·         liquid SOC or LB (0.5ml) (without Ampicillin.  Aliquot so students do not contaminate or omit step)

·         T4 DNA Ligase and buffer (1µl/group) (NEB)

·       pML104 or similar  digested plasmid (SmiI and BclI digested), pML104 undigested or pUC19 or other (+) control

·         cloning oligonucleotides

5’ phosphates are not required. Can order standard oligos

·         42C water bath

·         TE (Tris pH8 w/ EDTA) (1ml) for TA/instructor

Instructor/TA:

Instructor/TA: Prepare digested pML104 (do in large batch and can use until runs out, indefinite shelf life)

1. Plasmid prep (multiple mini-preps or maxiprep) pML104*

1. *plasmid must be prepped from  dam-/dcm- strain of E.coli or BclI restriction enzyme will NOT work

2. Digest pML104 with SmiI and BclI

1. 25ul of sterile water
2. 4ul 10X FD Green buffer
3. 8ul of 350 ng/ul pML104 (2.8µg)
4. 1.5ul SmiI (Thermo Fast Digest) (blunt digest)
5. Incubate *37C* for  30min (or 1h, 30C)
6.  +1ul BclI (Thermo FastDigest) (sticky end digest)
7. Incubate 37C for 30min (or 1h)
8. 80C for 10min to inactivate enzymes
9. Optional: Gel Purify via manufacturer kit.

1. BW has empirically determined that it is not necessary to gel purify.  Gel purification can reduce the number of colonies present when no insert is present, but without gel purification, 8/8 colonies PCR verified as containing insert when insert was present in ligation vs 0/8 without insert present, despite similar colony numbers on plates.

10. Store digested plasmid DNA at -20C.

3. Nanodrop to determine concentration
4. Test digested plasmid by ligating +/- insert and then colony PCR verify that insert is present.
5. Store digested plasmid at -20C

Instructor/TA: Prepare competent E.coli

1. Grow TOP10 (or similar) E. coli overnight on a shaker (LB no antibiotics!, use aseptic technique/flame carefully) in 2ml LB media
2. The next day, take a small volume of the overnight E.coli culture (~500 μL) and sub-culture it into another incubation flask containing 50 mL of LB.Grow at 37C with shaking, check the OD600 every 1 hours until it reaches OD 0.3 to 0.5.  (probably for 1.5-3hr total)
3. Put cells on ice. After this step put everything on ICE
4. Put the cells on ice for 10 mins (keep cold now on).
5. Collect the cells by centrifugation for 3 mins@5000rpm 4C
6. Decant supernatant and gently resuspend on 10 mL cold 0.1M CaCl2 (cells are susceptible to mechanical disruption, so treat them gently).
7. Incubate on ice x 20 mins
8. Centrifuge 3 mins@5000rpm 4C
9. Discard supernatant and gently resuspend in 5mL cold 0.1M CaCl2 + 15%Glycerol
10. Dispense in sterile 1.5ml tubes (100μL/tube) on ice. Freeze in -80°C.
11. Check the transformation efficiency of your chemically-competent cells by transforming with a plasmid that contains a positive selection marker (e.g., pUC19 & Amp). [BW verified that ~1.5 year old homemade competent TOP10 still transforms efficiently enough for this protocol to work.]

Instructor/TA: Prep LB+Amp or LB+Carb plates (at least 24h in advance)

Keep 100mg/ml Ampicillin or Carbinicillin stocks at -20C.  Plates are good for 1 month at 4C.

1. 10 g tryptone (or bacto peptone works fine)
2. 5 g yeast extract
3. 10 g NaCl
4. For 1L (makes ~40 plates)
5. FOR LBAmp AGAR plates - add ~1.7% agar (17g agar), [for liquid broth leave out agar]
6. Autoclave 121C 15PSI for 30minutes (time can depend on volume autoclaving)
7. let cool until doesn't burn you (but don’t let it solidify yet), then
8. Add 1000 µl of 100 mg/ml Ampicillin [final concentration in plates ends up 100 µg/ml] or Carbinicllin
9. Pour plates at ~25ml/plate.  Let solidify and store inverted at 4C in sealed bag or tupperware.
10. Plates are OK to use for about 4 weeks after made if stored at 4C

Instructor/TA: Prepare oligonucleotides as dsDNA – resuspend, combine, and anneal

1.        Briefly centrifuge the oligonucleotide tubes to make sure the material is at the bottom

2.        Dissolve each oligonucleotide in TE to make 100µM and mix by pipetting or vortexing

a.        Take the nmol amount on the tube and add 10times the amount of µl of TE

b.        e.g., if 28.4 nmol then add 284 µl of TE to the tube to make 100µM

3.        Make 1µM of the cloning oligos combined

a.        dilute 1µl of each of the two 100µM cloning oligos into 98µl sterile water in a pcr tube

b.                (be sure that you added 1µl of the second cloning oligo)

4.        Make a 50µM stock of the repair template oligos

a.        mix 50µl of both of the 100µM repair template oligo in a pcr tube

b.                (be sure that you added 50µl of the second repair template oligo)

5.        Heat oligo mixtures (2 tubes - one tube with both cloning and one tube with both repair template oligos) to 95C ~10min and allow to slowly cool to room temp for 30min (use Protocol on PCR machine ‘95DOWN’ )

6. store stock tubes and diluted dsOligos at -20C

Students

Ligate the annealed cloning oligos into the digested pML104 plasmid

1.  Setup the ligation reactions (two).

1. One tube will be with the insert (cloning oligos),
2. the other will be a ‘no insert’ (no oligos added) control.

1. X µl of digested pML104 (~10-20ng) [TA/instructor will provide volume to add]
2. 1 µl of 10X DNA ligase buffer
3. 1 µl of T4 DNA Ligase (NEB) (400 U/µl)
4. 1 µl of cloning oligo mix (1µM)  OR
5.           or 0µl oligos for ‘no insert control’
6. Y µl of sterile water (to 10µl total volume)

2. Incubate ligation mix for 1 hour at room temperature

F21 - add 1ul cloning oligos to mastermix, incubate 45-60min room temp

E.coli transformation

Transformation Samples (3): Ligation mix with insert, no insert control, and positive transformation control e.g., pUC19 or pML104 undigested.

1. thaw on competent E.coli (3 tubes) 10-30min on ice
2. Add plasmid DNA to be transformed

1. Into one labeled E.coli tube add 10 µl ligation reaction
2. Into another labeled E.coli tube add no insert control
3. Into another tube add 1µl of positive control plasmid DNA

3. Flick tube gently to mix
4. Put tubes into 42C water bath for 30sec
5. Put tubes on ice 20-30 minutes
6.  [can skip this step] Add 150µl SOC or LB (sterile)
7. Add 150µl onto one LB+100µg/ml Carbinicillin plate and disperse on plate using sterile glass beads
8. Incubate plate 37C overnight
9. Next day TAs will move plate to 4C fridge for next week

Lab 2 E.coli Colony PCR - verification of clones.

Background

        PCR background video 

Using one of the oligonucleotides you used to clone as a primer and the other primer internal to the plasmid, we will perform PCR to verify the presence of your cloned insert (gRNA sequence). We will expect a band of ~400bp from the PCR reaction if the insert is present in the plasmid (see the figure below).

Colony PCR technique 

Equipment and Materials (per group):

·         Apex Red 2X PCR Master mix (Genesee Scientific)

[contains Taq DNA polymerase, MgCl2, dNTPs, buffer, & loading dye]

·         One of cloning oligonucleotides and and M13R aka oBW626 primer (TTTCACACAGGAAACAGCTATGAC)

Oligos custom standard from ThermoFisher Scientific

·         Thermocycler

·         LB+Amp plates (1% tryptone, 0.5% yeast extract, 1% NaCl, 2% agar, 100 µg/ml ampicillin) Carbenicillin (CARB) can be used in place of Amp (plate shelf life ~1-3 months?)

Instructor/TA:

LB+Amp plates made last lab OK, if enough (plates are good for ~1month).

one of the specific cloning oligos at 10 µm (typically the 2nd oligo that starts with CTAG)

[does NOT start with GAT]

M13R primer (oBW626, tttcacacaggaaacagctatgac) diluted to 10µM.

-^[a]

Students

Count colonies on LB+Amp plates and record

Use a marker to put a dot on each colony as you count it

Count each plate and record your counts in your lab notebook

Count the positive, negative control, and experimental plates

        If too many in positive control, can score as many or >100

Polymerase Chain Reaction (PCR)

Colony PCR technique

Use 4 colonies from your cloning (+insert) plate and 1 colony from your no insert plate (as a negative control).  Keep track of which is which.

1.       Make a PCR master mix (in PCR tubes)

1. + 22 µl water
2. + 22 µl Apex Red 2X Master Mix
3. + 1 µl 10µM your 2nd cloning oligo *(oligo that starts with CTAG)*
4. + 1 µl 10µM M13R (oBW626) sequence: TTTCACACAGGAAACAGCTATGAC

2.       Add 10 µl of mastermix into each PCR tube (4total)

3.       Transfer some of a single E.coli colony to a fresh LB+Amp plate and label with e, Initials and unique number for each colony eBW1, eBW2, then put the pipette tip directly into the corresponding PCR tube (each PCR tube will have a different colony in it)

make it so that PCR tube 1 = eBW1, etc.

4. Put into thermocycler (PCR Machine) and TA will run PCR ‘CRISPRCOL’ (takes ~1.5h)

o   95C 3min for 1cycle

95C 30sec

 52C 30sec

 72C 35sec

             For 32 cycles

o   72C 3min for 1 cycle

o   12C infinite (hold at 12C)

Lab 3 Gel electrophoresis of E.coli colony PCR and Plasmid prep.

Brief background video about gel electrophoresis

Technique video (pouring, running and visualizing a gel)

Equipment and Materials (per group):

Agarose,

50X TAE buffer (to make 1X TAE to be used).

TA: For 50X TAE - On top fridge in green box in biochem lab.  

If needed to make, add 242g/L Tris Base, 18.6g/L, 57ml /L Acetic acid* (*use PPE and fume hood with another person present)

10mg/ml Ethidium Bromide (EtBr) -  

Apex Safe Stain (5 µl per 100ml of gel); (UV 290nM or blue light 490nM excitation and emission is at ~520&635nM)

can use SybrSafe protocol on geldoc

Gel electrophoresis rig

PPE (gloves,lab coat )

UV/blue gel doc imager

GeneRuler 1kb Plus DNA ladder

Gel electrophoresis

Instructor/TA:

Pour gels (video)

Per each gel make 1% (1g agarose per 100ml 1X TAE buffer).  Volume depends on gel apparatus size.  

1. Dissolve the appropriate amount of agarose into the appropriate amount of  TAE.  Microwave until just starting to boil and immediately stop the microwave. Use Caution to avoid boiling over when touching.
2. After cooled for ~5minutes, add 10µl of Ethidium bromide per 100ml of buffer.  Use PPE when working with Ethidium bromide. add 5ul Apex Safe Gel Stain per 100ml gel
3. Pour gel into casting tray, add comb(s) and allow to gel to solidify

Start O/Ns for plasmid preps

1. Sterilly inoculate student E.coli into 2-3ml LB+Amp (or LB+Carb) liquid and shake 37°C overnight.  Label student samples appropriately. Innoculate at least 2 colonies per student.
2. Make aliquots for plasmid preps.  Be sure that Ethanol is added to the Kit Plasmid Wash Buffer before using.

Students:

Run Gel Electrophoresis

Pouring, running, and visualizing a gel videos

Load Samples into gel

1. Load 7µl of a DNA ladder standard
2. Load 10µl of your PCR reactions each into a separate well

1. Record location of where you loaded your samples

Run Gel

1. Run gel at ~125V for 30-60minutes.

Image Gel

1. Using GelDoc instrument in Biotech lab
2. video instructions for GelDoc

Plasmid Prep (Zymo ZR Plasmid miniprep kit)

(May vary if we purchase from a different manufacturer or if we DIY in future, or could use boiling lysis prep)

1. Spin 1.5ml of E.coli overnight culture for 20sec in microcentrifuge (2 tubes from 2 overnights).
2. Discard supernatant (liquid above pellet)
3. Resuspend pellet in 200µl P1 buffer (RED) by pipetting
4. Add 200µl of buffer P2 (Green), mix by inverting 2-4 times. Cells are completely lysed when the solution appears clear, purple, and viscous. Proceed to the next step within 1-2 minutes.
5. Add 400 µl of P3 Buffer (Yellow) and mix gently but thoroughly. Do not vortex. The sample will turn yellow when the neutralization is complete . Allow the lysate to incubate at room temperature for 1-2 minutes.
6. Centrifuge samples for 2 minutes.
7. Place a Zymo-Spin IIN column in a Collection Tube and transfer the supernatant from Step 5 into the Zymo-Spin IIN column. When pipeting the supernatant be careful not to disturb the green pellet to avoid transferring any cellular debris to the column.
8. Centrifuge the Zymo-Spin IIN/Collection Tube assembly for 30 seconds
9. Discard the flow-through in the Collection Tube, making sure the flow-through does not touch the bottom of the column. Return the Zymo-Spin IIN column to the Collection Tube.
10. Add 200 µl of Endo-Wash Buffer to the column and centrifuge for 30 seconds.
11. Add 400 µl of Plasmid Wash Buffer to the column. Centrifuge for 1 minute
12. Transfer the column into a clean 1.5 ml microcentrifuge tube and then add 30 µl of DNA Elution Buffer to the column. Centrifuge for 30 seconds to elute the plasmid DNA.
13. With TA, use the nanodrop to determine concentration and quality of plasmid DNA (there should be a clean peak centered near 260nM without large amount of absorbance below that) Record concentration on tube and in lab notebook. Save Abs vs wavelength output from spec and include in lab notebook. OK to just take a screenshot with smartphone or email image/file.

Nanodrop video

If not good, can run through PCR cleanup kit or EtOH ppt (as long as some abs at 260 is present)

14. Optional: Send plasmid for whole plasmid sequencing (e.g., send 30ng/ul to Plasmidsuarus for $15/plasmid as of 1/2022).

Lab 4 Yeast transformation

Brief background video:

Basic techniques:

Innoculating a yeast overnight

Patching and streaking yeast strains

Aseptic technique*

Equipment and Materials (per group):

CSM-URA plates (3 plates per group) with YNB+nitrogen and 2%glucose

Zymo EZ Yeast Transformation kit (or see Geitz et al. for DIY PEG based transformations)

pho8 yeast or BY4742 wildtype (Su21&F21 we used pho8 knockout yeast strain)

repair template oligonucleotides (50µM dsDNA) previously ordered

Experimental student crispr plasmid that was cloned and verified

pML104 plasmid (NOT digested) or other URA3+ positive transformation control plasmid (e.g., pRS425) ,

30C incubator

Instructor/TA: Preparing frozen yeast competent cells

1. Start overnight of WT BY4742 or pho8 mutant

1. [*or just inoculate single colony into 1L YPD 30C shake and check OD next day for log phase*]

2. Next morning, dilute 10ml overnight into 1L YPD and grow yeast cells to log phase (ideally OD 0.8-1.0) at 30C with shaking
3. Spin down cells, discard supernatant (spnt) (this is liquid above pellet, leave pellet)
4. Resuspend pellet in 20ml Zymo EZ Solution #1
5. Spin down cells, discard spnt
6. Resuspend in 12ml Zymo EZ Sol #2.  Then aliquot 50ul per 1.5ml tube (4 sections x20 students x3 transformations  = 240tubes x 50µl = ~12ml)
7. Freeze wrapped in paper towels in closed styrofoam container at -80C
8. Competent yeast cells are likely good for 1 year, TBD.
9. TA test yeast competency (before deploying with students) by transforming a positive control plasmid (e.g., pML104).

Students: Yeast Transformation Protocol:

Three Transformation tubes:  #1 control plasmid (pML104). #2 Experimental plasmid only, and #3 Experimental Plasmid + repair template (‘RT’)

1. Mix 50 µl of competent cells with 0.2-1 µg plasmid DNA (in less than 5 µl volume);
2. To tube with repair template, also add 50ul of repair template (RT) oligos
3. add 500 µl ‘Frozen-EZ Yeast Solution 3’ and mix thoroughly by pipetting up and down.
4.  Incubate at 30°C for 45-60 minutes. Mix vigorously by flicking with finger or vortexing 2-3 times during this incubation
5. Spin down cells 30sec at ~5000rpm in microcentrifuge (be sure to appropriately balance tubes)
6. Discard 250µl of supernatant liquid above pellet, leave remaining supernatant liquid
7. Resuspend cell pellet in the remaining volume and spread all the remaining volume of each transformation on separate labeled -URA plates (label with initials and transformation description - e.g., pos control, exp plasmid, exp plasmid + RT). Spread using sterile glass beads.
8. Recycle glass beads into collection beaker (TA will ethanol rinse, dry, and then autoclave)
9. Incubate the plates at 30 °C for ~3-5 days to allow for growth of transformants (incubate inverted, agar at top and lid at bottom to prevent condensation from dripping from lid to agar). Store 4 °C if needed.

Lab 5 Yeast colony PCR^[b][c][d]

Background: See PCR lab 2 for background on colony PCR.

PCR Primers used:  oBW27 GCCTTATAGCTTGCCCTGAC and oBW28 ACCCCTAGATTTTGCATTGCTC are just upstream (5’) and downstream (3’) of the PHO13 gene and should amplify a ~1 kb product consisting of the PHO13 gene.

Equipment and Materials (per group):

o27 and o28 primers (diluted at 10µM each and add equal volumes of each to a tube to make 5uM of both). 2x Apex Red PCR mix (Genesee Scientific).  Freshly grown yeast (not stored at 4C over weekend, repatch day before use) from Lab 4. Ladder=GeneRuler 1Kb Plus

Instructor/TA: Prepare yeast and primer mastermix

If plates have been stored at 4C, freshly patch 4 colonies from the transformation plate onto a -URA plate the day prior for colony PCR and incubate at 30C overnight. Make sure freshly grown yeast are used in yeast colony PCR reactions.

Students:

Count and record the number of colonies on your plates (if too many to count on positive control plate, you can just record that plate as  ‘hundreds’)

Yeast Colony PCR

1. Make PCR Mastermix:

125µl 2X Apex Red

125µl water (autoclaved dH2O)

6.3µl 10uM  primer mix (o27 and o28)

2. Aliquot 50µl into 4 PCR tubes
3. Label your PCR tubes (include your initials).  

1. Be sure you know and record which yeast colony is in which tube.

4. Touch each yeast strain with a sterile pipette tip and transfer into a PCR tube with mastermix
5. Run PCR reaction: (PCR protocol PHO13AMP; thermocycler takes ~2.5hr)

1. 95C 5min
2. 95C 30s
3. 55C 30s
4. 72C 60sec

Repeat steps 2-4 for 32 cycles

5. 72C 5min

• *Test fast pcr so can do more in lab session.

Lab 6 Gel electrophoresis of yeast colony PCR, PCR Clean up, Sequencing

Equipment and Materials (per group):

PCR Clean up kit^[h] (ThermoFisher GeneJet PCR Cleanup kit) [or EtOH and 3M Na Acetate].  The kit will remove loading dye and PCR primers from the PCR product.

Gel Rig, Ethidium Bromide or Apex GelSafe stain, Agarose, ladder, (see lab 3)

Sequencing oligos (or could a PCR oligo):

Instructor/TA: Prep gels beforehand. Be sure to Add EtOH to kit Wash buffer before 1st use.

Students: Protocol

1. Run Gel Electrophoresis (gel electrophoresis technique videos)

1. Load 7µl of a DNA ladder standard
2. Load 10µl of your PCR reactions (each into a separate well)
3. Run gel at ~125V for 30-60minutes.
4. Image gel using GelDoc XR instrument in Biotech lab

2. PCR Cleanup (kit instructions)

1. Add 40 µl Binding Buffer to 40µl PCR mix (1:1 ratio). Mix thoroughly by pipetting.
2. Transfer the solution from step 2.1 to the GeneJET purification column.
3. Centrifuge for 30-60 s (~full speed in microcentrifuge). Discard the flow-through liquid that passes through the column. Save the column that now contains your bound DNA.
4. Add 400 µL of Wash Buffer to the GeneJET purification column.
5. Centrifuge for 30-60 s. Discard the flow-through liquid that passes through the column and place the purification column back into the collection tube.
6. Add 200 µL of Wash Buffer to the GeneJET purification column. Discard the flow-through liquid that passes through the column and place the purification column back into the collection tube.
7. Centrifuge the empty GeneJET purification column for an additional 1 min to completely remove any residual wash buffer. Note. This step is essential as the presence of residual ethanol in the DNA sample may inhibit subsequent reactions.
8. Transfer the GeneJET purification column to a clean 1.5 mL microcentrifuge tube. Add 25 µL of Elution Buffer to the center of the GeneJET purification column membrane (but do not touch pipette tip to membrane) and centrifuge for 1 min.
9. Discard the GeneJET purification column and store the purified DNA at -20 °C.

3. Nanodrop. nanodrop use video

1. Clean nanodrop gently with wet kimwipe and gently dry with kimwipe
2. Add 1ul elution buffer as blank, blank machine, dry block
3. Add 1ul sample, read sample, save spec and record concentrations

4. Send for sequencing

1. To the sequencing tube, add 5µl of 40-60ng/ul (200-300ng) cleaned up colony PCR product and 5µl of 10µM sequencing primer.  BIOL4242 will use Eurofins Genomics sequencing service.  The first ~20nt of sequencing data may not be good.  Pick a sequencing primer that is not too close (<25bp), but not too far (ideally within ~500bp) of your mutated region of Pho13). Can also use o27 (same as PCR primer before gene start) to sequence.  o28 has been hit or miss (less consistently got good seq data, so maybe use o10 instead).
2. Alternatively: use amplicon sequencing instead of Sanger and no sequencing primer is necessary). Check company recommendations for sequencing prep for their amplicon sequencing service.

Available sequencing primers:

• Replace Sanger with amplicon using next gen - more expensive but less failure and better read depth and opportunity to teach about newer tech

Students Post Lab: Sequence analysis

1. Tutorial video 
2. Open the .abi file with FinchTV
3. If peaks are OK, copy and paste sequence into ApE

OK                                                   NOT OK

4. Also open your repair template sequence and Pho13 nucleotide sequence in ApE windows
5. In ApE click ‘Tools’ ‘Align sequences’ and verify the presence of your desired mutations in the sequencing data.

Lab 7 Lysis & Enzyme Assays^[i] and BCA

Background^[j][k]:

Pho13 is an alkaline phosphatase enzyme that can use paranitrophenyl phosphate (PNPP) as a substrate. You made an amino acid substitution to the Pho13 enzyme.  PNPP is clear when dissolved in aqueous solutions and can be hydrolyzed to PNP by alkaline phosphatases such as Pho13.  Para nitrophenol (PNP) is the product of the enzyme reaction and is being used to generate a standard curve for the concentration of PNP with an absorbance maximum at 400nM. We may perform experiments in a yeast strain where another alkaline phosphatase encoding gene (PHO8) is removed from the genome in order to simplify the system and have only a single enzyme with alkaline phosphatase activity present. We will lyse cells using Cell Lytic Y, a proprietary commercial detergent based lysis solution^[l].

Basic techniques

Patching and streaking yeast strains

Equipment and Materials (per group): Cell Lytic Y (Sigma), BCA kit (kit has 2mg/ml BSA std), Yeast cells grown overnight in YPD media (1% yeast extract, 2% peptone, autoclave, 2% glucose), PNPP, PNP, 20mM Tris pH 9.0 + 20mM MgCl2 buffer, 96 well plates (wash well w/DI water after finished, dry, and reuse 96 well plates)

Instructor/TA prep:

1. Make 10mM PNPP in 20mM Tris ^[m]pH9.0 + 20mM MgCl2  (make 2x 1.1 ml/student)

1. Dissolve 0.372g PNPP (disodium salt hexahydrate) into 100ml buffer.
2. Make week of experiments to be safe for now.  Shelf life is long in water, but unknown in buffer w/Mg (probably OK, TBD). After ~6months stock in water turned slight yellow (clear when first dissolved).
3. Store in dark container at 4C.

2. 20mM Tris pH9.0 + 20mM MgCl2 buffer (10ml/student)
3. Make 0.5mM PNP standard (1ml/student)
4. Mix BCA solution just before class starts according to mfg instructions (50:1 reagent A/B) (~5ml per student). Aliquot BSA standard (30 µl of 2mg/ml per student)
5. Specifics TBD - Day before, inoculate a single yeast colony (or can use more cells from large patch of cells to be sure O/N is grown densely for next day) into 30ml YPD per group (for each control pho8 and mutant strain) and make a 1/10 and 1/100 dilution into 40ml YPD., could make larger volume O/N and aliquot when applicable.
6. Shake 30C overnight (~12h) and students use sample the next day, OD600 = ~ 0.7-1.0 (Log phase cells).
7. After students are done with experiments, rinse out 96 well plates well with distilled water and let dry (upside down) and then the plates can be reused for future assays

Students:

Lysis

1. You will lyse: Wildtype (BY4742 or “WT”), pho8 deletion mutant, pho13 deletion mutant, and your mutant (4 total samples). Fall2021 your mutant was made within a pho8▲ deletion strain and you will use a control pho8▲ (BW1418) cells and either pho8 Pho13-G253A or (BW1423) OR pho8 Pho13-D196G (BW1426) strains.
2. Samples from TA - Spin down yeast culture in 2000rpm for 5min, pour off and discard the supernatant (liquid above pellet)
3.  Resuspend the cell pellet in 500 µl of Cell Lytic Y, mix by pipetting up and down, and transfer the suspension to a 1.5ml or 2ml tube
4.  incubate for 30 min at room temp (with agitation)

1. While sample is lysing for 30min, you can: (See detailed steps on following pages)

1. pipette BCA sol. into microplate wells
2. serial dilute BSA protein standard in tubes
3. Serial dilute PNP standard in microplate
4. Serial dilute 10mM PNPP in tubes to make 5mM-0.31mM PNPP solutions
5. Pipette PNPP into microplate
6. *Don’t forget to do remaining Lysis steps 5-6 after the 30min incubation before using in the BCA and enzyme assays

5. Spin 13,000xg in microcentrifuge for 10min
6. Transfer ~500µl supernatant (liquid above pellet) to fresh tube (try not to do not disturb pellet, which is unlysed cells and cell debris). The clear transferred supernatant is your cell lysate that you will use for BCA and enzyme assays.
7. Proceed to BCA and then enzyme assays below
8. After the lab is completed, store protein at -20C (initial & label frozen lysate tubes)

BCA Protein Concentration Assay

1. TA/instructor: Mix BCA reagent A and B (50:1) as indicated by manufacturer. = BCAsol
2. Add 145µl of BCAsolution to wells indicated above (E1-E3,F1-F3, G1-G12 and H1-H12)
3. In 1.5ml or 2ml tubes, serially dilute 20 µl of 2mg/ml BSA protein standard ½ (so 20µl 2mg/ml BSA into 20µl of water, then 20  µl of 1mg/ml into 20ul water to make 0.5mg/ml, etc) to make 1, 0.5, 0.25, 0.125 mg/ml BSA. (see image)
4. Serially dilute ½ pho8 lyaste, mutant lystate, and cell lytic Y by diluting 20µl into 20µl buffer and then take 20µl of the ½ dilution into 20µl of buffer to make ¼
5. Try to perform the following steps quickly and in succession. See also the plate map above.

1. Add lysate to wells around the same time as standard is added to wells so they get roughly equal incubation times

6. Add 5µl of cell lytic Y to E1 and F1; ½ cell lytic Y to E2 and F2, ¼ cell lytic Y to E3,F3
7. Add 5 µl^[n] of

1. 2 mg/ml BSA to G1 and H1 (see plate map above for well locations)
2. 1 mg/ml BSA to G2 and H2;
3. 0.5 mg/ml BSA to G3 and H3
4. 0.25 mg/ml BSA to G4 and H4;
5. 0.125 mg/ml BSA to G5 and H5

8. Leave G6 and H6 blank (zero BSA added)
9. Add 5µl of pho8 lysate to wells G7, H7
10. Add 5µl of ½ pho8 lysate to wells G8, H8
11. Add 5µl of ¼ pho8 lysate to wells G9, H9
12. Add 5µl of mutant lysate to wells G10, H10
13. Add 5µl of ½ mutant lysate to wells G11, H11
14. Add 5µl of ¼  mutant lysate to wells G12, H12
15. Incubate at room temperature

1. While incubating, setup Enzyme Assays below

16. Read Absorbance at 562nm on microplate reader

Enzyme Assays

Make PNP standard curve

1. Carefully, add 300µl of 500µM PNP standard to wells A1, B1, C1 (See platemap below)
2. Add 150ul of buffer (Tris pH9 +20mM MgCl2) to wells A2-A6 and B2-B6 and C2-C6
3. In the microplate, serially dilute 150ul from A1 to A2, then from A2 to A3, A3 to A4, A4 to A5 (discard remaining 150µl)

1. leave A6 without any PNP

4. Repeat serial dilution using 150ul from B1 to B2, and serial dilute to B5 and repeat again from C1 to C5 (leave A6,B6,C6 as ‘0’ or ‘blanks’ without any PNP)

Serial Dilute Substrate (PNPP)

5. Serial dilute PNPP substrate 1/2

1. In 2ml tubes, serially dilute pipette 1000µl of 20mM PNPP (in buffer)  into 1000 µl buffer (20mM Tris pH9+20mM MgCl2) to make 10mM, 5, 2.5, 1.25, and 0.6mM  PNPP solutions

Perform Enzyme assays

6. Add 145 µl of the indicated concentrations of PNPP  to wells as shown in the plate map below (e.g, 20mM ^[o]goes into A7-F7 column & D1; 10mM into A8-F8& D2, column,etc.)
7. Try to perform the following steps in a timely succession.  Start timing at the start of the next step.
8. Add 5µl of cell lytic Y to wells D1-D6 (blanks)
9. Add 5µl of pho8 lysate to wells A7-A12 row, B7-B12, C7-B12
10. Add 5µl of your ‘mutant’ lysate to wells  D7-F12
11. Time and Incubate for 10 mins at room temperature [if no yellow observed yet in A7-F7, wait longer 20-30min]
12. *Wearing Safety Glasses and Gloves*: Stop Enzyme reactions by adding 2 µl^[p][q][r] of 2M NaOH

1. (STOP in same order as started to get each reaction the same time incubation)

13. Read Absorbance at 400nM on microplate reader (also read BSA assay 562nM at same time)

Lab 8x Chromatography: DEAE (anion exchange) omit

Youtube link

Background

Column chromatography is used to purify proteins and allows for retention of enzymes in the folded active state. We will use Diethylaminoethyl cellulose (DEAE cellulose), an anion exchange resin to perform ion exchange chromatography. Note that the pKa of the DEAE amine group is 10, so the pH of the chromatography experiment should be below that to ensure the resin is positively charged. The isoelectric point (pI) of the yeast Pho13 protein is ~6, so above that pH the protein will carry a negative charge.  During chromatography, we will bind our protein to the DEAE column, then wash the column, then elute protein from the column.  In this experiment, the elution will be performed using NaCl.

Structure of DEAE: (Cellulose is a polysaccharide consisting of many β(1→4) linked D-glucose)

Equipment and Materials (per group):

Reagents

DEAE Cellulose,

NaOH, HCl, NaCl,

20mM Tris pH 9,

p20, p200 pipettes, 0.5ml tubes 1.5ml tubes

10ml disposable columns or similar (TA clean and reuse columns),

BCA kit, 2mg/ml BSA standard

PNPP (10mM or 1mM?)

Recipes: 10X Tris (200mM): dissolve Xg of Tris base into less than 1 L water and adjust pH to 9.0 with HCl then bring volume up to 1L.  Then dilute to make 20mM (100ml 10X into 900ml water).

Instructor/TA:

This lab worked well with wildtype, but not very well with pho8 mutant (low activity observed).  Growth conditions to maintain Pho13 expression might need to be determined (Log phase vs stationary phase cells?). Elution conditions (elution volumes?) could be optimized.

Prep BCA solution right before lab or give appropriate aliquots BCA A and BCA B solution.

Make PNPP stock within ~2 months of use and store in dark container at 4C^[s][t][u].

Ice for students

DEAE Preparation

1.       suspend dry DEAE cellulose resin in 5 volumes diH2O and allow to swell and settle for 30-45 minutes.  (students will need ~5ml swollen resin per column)

2.       Measure volume of settled resin to determine Column Volume (CV) for washing steps.

3.       Filter suspension and re-suspend resin in 2 CV of 0.1M NaOH+0.5M NaCl for 10 minutes.  (or can load onto column and allow different liquids to sit on column for 10min)

4.       Pour slurry into Buchner funnel and allow buffer to slowly drain off (~1 V buffer in 5 minutes).

5.       Repeat using 0.5M NaCl (without 0.1M NaOH)

6.       Repeat using 0.1M HCl+0.5 M NaCl

7.       Repeat using diH2O

8.       Repeat using 20mM Tris pH 8.1.  Could check to make sure pH is correct coming off column.

9.       Load ~3ml DEAE resin into column with cotton plug, wash with some buffer

10.   After students have finished, collect all resin and redo NaOH/NaCl/HCl washes as above, then resuspend the resin in 2 CV of 1M NaCl and adjust pH to 7-8 with NaOH and store at 4°C.

Students

DEAE chromatography ^[v]

*Keep the column from drying out - should always have liquid above the column resin*.

1. Remove 35ul of Lysate and store for future (ice).
2. Run buffer to just into column matrix and load all of yeast lysate plus 1.5ml of Tris pH9 buffer onto column.
3. Wash column with 3ml buffer (2xCV, assuming CV=3ml: CV=column volume) and collect entire volume as ‘flow through’.
4. Let buffer run to top of DEAE gel bed, then slowly add 1ml buffer+0.1M NaCl then immediately begin collecting 0.5ml fractions^[w]
5. Let buffer run to top gel bed, Add 1ml buffer + 0.3M NaCl
6. Let buffer run to top gel bed, Add 1ml buffer + 0.6M NaCl
7. Let buffer run to top gel bed, Add 3ml buffer (without NaCl)
8. Once buffer has run to top of gel bed can stop collecting fractions
9. Add 4ml buffer and allow to flow through^[x]
10. Load next lysate and repeat (starting at step 2)
11. Use fractions for BCA and Enzyme assays

BCA protein concentration assay on fractions.

*TA: Mix 50:1 BCA reagent A to reagent B to get enough volume for students

1.  Load 140µl of BCA reagent for enough wells for fractions and for standard curve and blank.

2. Serial dilute 2mg/ml BSA standard ½ in water (30 µl into 30 µl water).

3. Load 10µl of fraction or BSA standard to appropriate wells

4. incubate 10-30min room temp

5. Measure absorbance at 562nM

Can store fractions in freezer at -20C

Plot Enzyme assay and protein on same graph (can just plot absorbances for now to be quick, but save data)

Enzyme assays on DEAE fractions

1.       Load wells A1-A8 with 150 µl Tris buffer.  You determine where on the plate and record in your notebook.

2.       Add 150ul of 1mM PNP standard (yellow) to A1 and A7 wells with 150ul of Tris buffer and mix to make 500µM PNP in first standard well, then remove 150ul of this to next well containing 150ul of buffer and mix and continue serial dilute ½ for standard.  Include a 150µl buffer only as a blank (A6&A12)

3.       Load 140ul of 1mM PNPP (substrate) in 20mM Tris pH8.1 + 10mM MgCl2 into microplate in enough wells for all fractions

4.       Add 10 µl of each fraction to microplate that has 140ul PNPP already added

5.       Let incubate for 10 minutes

6.       Read Absorbance at 400nM (_4242AlkalinePhosphatase protocol)

X Lab X SDS-PAGE of Fractions

Background be added later

how to setup and run gel rig https://bit.ly/317kYAp

Objectives:

·         To empirically prepare, load, and run enzyme fractions on an SDSPAGE gel

Experiments:

·         Given your fractions concentrations, determine the volume needed to load a specific quantity of each fraction

·         Load, run, and visualize F1,F2, F3,... on an SDSPAGE gel

Reagents:

TA Prep work:

**Be comfortable with how to setup and run gel rig** https://bit.ly/317kYAp

Set heat blocks to 90C

Make 1X SDS PAGE running buffer from 10X

With students, remove gels from plastic wrap and setup gel rigs (illustrate process to students)

* be sure to remove green strip from bottom of gel*

To be performed by Students:

Preparation Protein sample for loading

1.       Calculate the amount of sample needed to load 15 µg of protein on a gel

2.       Make Fraction samples to load in gel, which will be:

2X Laemmi                                  15 µl

Fraction  (X µg/µl)                 X µl

Water                                           (15 – X) µl

Total                            30µl

3.       Heat samples at 90C for 5min

4.       Load 10µl of each sample into the well of a gel.

Pipette slowly and carefully, TA will assist with loading.

Record the gel # and well positions that you are loading into

Make note that we’re loading 10µl/30µl or 1/3 of sample (e.g., 5µg was loaded instead of 15µg), in part to give more chances in case loading errors occur.

 Example calculation:

If F1 was 1.5 µg/µl and you wanted to load 15µg total of BSA protein

So, to have 15  of BSA ready to be loaded into the gel:

F1 protein (1.5 )              10

2X Laemmi                                    15µl

Water                                              5µl

X Lab X other assays

Background

Allow for repeat experiment time and/or allow for students to choose additional experiments and provide some example possibilities

Equipment and Materials (per group):

 Enzyme assays?

-Test range of pH buffers: 7.0, 7.5, 8.0, 8.5, 9.0 (Tris base can work across this range)

        Have students make own buffers

-Test a range of MgCl2 concentrations: 50, 20, 10, 5, 1, 0.5, 0 mM

-Test different temperatures: room temp, 30C, 37, 50, 65°C using heating blocks

-Inhibitors; inorganic phosphate, ZnCl2,  student hypothesis to test given an inventory of chemicals?

Yeast assays

growth rate using platereader, any known pho13 null phenotypes?

sensitivities plate/liquid growth rate,

student hypotheses?

Additional possible experiments

Pho8-GFP or tag w/HA

western blot, - dot blot?

microscopy

Could measure GFP fluor in fractions?

X Lab X Sephadex gel filtration (size exclusion) chromatography

CURRENTLY NOT IN USE AND INCOMPLETE

Background

Equipment and Materials (per group):

Let Sephadex G200 swell in Tris pH8 buffer overnight (maybe we should get G100 for future)

Load ~3ml sephadex into column

Sephadex gel filtration chromatography

1.       Remove 100ul aliquot of pooled fraction from DEAE purification and store at -20C for future use

2.       ?Add 100ul of 5mg/ml Dextran Blue to pooled fractions

3.       Let buffer run to gel bed, then load fraction onto column, let liquid run into gel bed, add Tris buffer pH8? carefully and do not disturb gel bed (and don’t let gel bed dry out as continuing, add more buffer as needed)

4.       Collect buffer in a clean beaker until dextran blue is about to come off, then measure volume of collected buffer that flowed through column and record as Void Volume (VO)

5.       Begin collecting ~0.3 ml fractions as dextran blue elutes from column, for (~24 fractions?)

6.       Transfer fractions to ice as they are collected

Enzyme assays on fractions

1.       Load 8 wells with 150ul Tris buffer.

2.       Add 150ul of 1mM PNP (yellow) to first well with 150ul of Tris buffer and mix, then remove 150ul of this to next well and mix and continue serial dilute ½ for standard.  Include a 150ul buffer only Blank

3.       Load 140ul of 1mM PNPP in 20mM Tris pH8.1 + 10mM MgCl2 into microplate in enough wells for all fractions

4.       Add 10 µl of each fraction to microplate that has 140ul PNPP already added

5.       Let incubate for 10 minutes

6.       ??Stop reaction with Xul 20 mM NaOH (ice-cold)

7.       Read Absorbance at 400nM (_4242AlkalinePhosphatase protocol)

Run BCA assay on fractions.

TA: Mix 50:1 BCA reagent A to reagent B to get enough volume for students

1.  Load 140ul of BCA reagent for enough wells for fractions and 8 wells for standard curve and 1 well for blank.

2. Serial dilute 2mg/ml BSA standard ½ in 8 tubes

3. Load 10ul of fraction or BSA standard to wells

4. incubate 30min room temp or 37C if available

5. Measure absorbance at 562nM

write up:

Using fraction with highest specific activity following sephadex,

-Test varying substrate concentrations

10mM, 1mM, 0.5mM, 0.25, 0.125 mM PNPP

Determine Vmax, Km, Kcat

X EtOH ppt- Ethanol Precipitation to Clean up yeast colony PCR prior to sequencing (Alternative to PCR clean up kit)

1. Add 1/10 volume 3M Sodium Acetate (pH ~5)
2. Add 2.5x volumes of 95%+ Ethanol (or can use 1 volume 100% isopropanol)
3. Optional: Leave in -20C for 15minutes or more
4. Spin down in microcentrifuge full speed for 10min (at 4C if possible)

1. Be careful not to disturb pellet from this point on
2. Sometimes pellet may not be visible (assume pellet at bottom/back of tube - back being side of tube facing outside of centrifuge)

5. Discard supernatant (can pour off or pipette off slowly/carefully)
6. Add  1ml of 70% ethanol (ice cold is ideal)

1. Spin down 5min full speed (if possible at 4C)
2. Discard supernatant (remove last bit with pipette, but don't remove pellet)

7. Let pellet dry at room temperature (could leave overnight open on lab bench with kimwipe covering opening to avoid dust) OR could use SpeedVac 10min spin w/vaccum
8. Resuspend in desired volume of TE or water (e.g., 50ul)
9. Nanodrop to determine concentration