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The objective of the lab course is to construct a new plasmid cloning vector called pTA7. This vector will be an E. coli / Saccharomyces cerevisiae shuttle vector. This vector will be useful for cloning using the Yeast Pathway Kit, see the next section for more details.

In the mec research group, we are interested in understanding and engineering the biosynthesis of fatty acids and related products by the unicellular fungi known as baker's yeast S. cerevisiae.

Genetic engineering of complex traits require the simultaneous deletion and/or expression of multiple genes. This is a challenging problem as genetic engineering is time consuming. To solve this problem, we developed a protocol for the assembly of metabolic pathways that we call the Yeast Pathway Kit (YPK). See our publication in ACS Synthetic Biology for more details.

We use this protocol for quick construction and expression of large metabolic pathways in baker's yeast Saccharomyces cerevisiae such as this heterologous fatty acid synthesis pathway. Plasmids such as the one we will construct in this lab course are used to propagate these constructs in E. coli or S. cerevisiae. In order to do this they need a selection marker for E. coli and one for S. cerevisiae. They also need an origin of replication for E. coli and one for S. cerevisiae.

The first plasmid we used for this purpose in YPK was called pYPKpw and it has the following functional parts (Table#1):

The Δcrp gene is inactive and only provide a recombination site. The pathways that we make are meant for S. cerevisiae, but we often need to transfer the pathway to E. coli so we can obtain larger amounts of higher quality DNA for analysis or transformation.

The pUC origin of replication (ORI) results in a high copy number of the vector in E. coli which is an advantage since it makes it easier to obtain large amounts of pure DNA. However, we have observed genetic instability in E. coli for some large pathways that we suspect is linked to high copy number. Our experience is that a lower copy number provides more stability.

We conceived a series of plasmid vectors called pTAx where x is a number from 1 to 11 (at the moment). The pTAx vectors were designed to have a lower copy number in E. coli to try to solve the stability problems of pYPKpw. The pTAx plasmids should have a lower copy number in E. coli than the pUC based pYPKpw since they have the pBR origin of replication that includes the ROP gene. The first pTAx plasmid, pTA1 was constructed by a former post-doc in the group, Tatiana Andrevna, hence the name.

The pTAx vectors are made from five genetic elements (Table #2), see (pTAx assembly strategy).

Each element is a distinct segment from a particular source plasmid. The pTAx vectors are similar to each other, but differ in the selection markers (yeast marker) and yeast origin of replication (yeast ORI).

The lab course is divided into nine practical classes. Each student attends three of the nine classes.

Take pictures during class! This helps remembering what you did and how. You can upload your pictures to this shared Google album. Use the link in Blackboard.

We will have a photo contest within each PL and we will also select the best image overall. This means that there will be up to four winners, one for each PL and one overall. Mark you image with your name, number and PL in the Google photo album.

material#LAB1

Summary:

• Plasmid miniprep using alkaline lysis

We will prepare four plasmids using alkaline lysis mini prep (see Table#3). These plasmids are the source for each of the five genetic elements. One plasmid is used as the source of two elements. The last column states why we need each particular plasmid.

We will use a homemade alkaline lysis plasmid miniprep protocol. Here is a short protocol for printing.

The teacher has prepared E. coli cultures in liquid or on solid medium beforehand.

Cultures with each plasmid were grown in or on LB with ampicillin for selection of the plasmids.

These videos show how to prepare plasmid DNA from E. coli using basically the same protocol.

material#LAB2

Summary:

• Gel#1 on plasmid DNA.
• Plasmid DNA dilution.
• Preparation of a PCR reaction.
• Make liquid YPD medium for LAB3.
• Make solid SD medium for LAB4.

1. Unfreeze the plasmids from LAB1
2. Vortex the tube to mix the content. If necessary, spin the tube for ~2-3 seconds to collect the liquid at the bottom.
3. Take out 10 µL of the content to a fresh tube.
4. Add 2 µL 6 x loading buffer to your plasmid DNA.
5. Put the gel in the gel tray in a square petri dish. The gel should have at least one well per group number and one extra for the marker.
6. Add 5 µL of the plasmid DNA to an empty well, start with the leftmost well.
7. All group members should load their plasmid.
8. Take note of where your samples are.
9. The teacher will help you to put the gel in the electrophoresis chamber.
10. The teacher will load the molecular weight marker.
11. Apply the electrical field as soon as you are done.
12. The electrophoresis last around ⌛15 min at ⚡200 volts in the Bachman gel tank using a homemade rectifier.
13. When the gel run is completed, the teacher with take a picture using a transilluminator.

• Put the gel in TAE + Midori Green, incubate 15-30 min
• Take picture

14. Pipette 1 mL (1000 µL) ultra-pure water into a clean 1.5 mL Eppendorf tube
15. Mark this tube well so that you can identify it.
16. Transfer 5 µL of the plasmid DNA to the tube with 1 mL water This will make a 5 µL/1000 µL = x 200 dilution.
17. Vortex the tube with the x 200 plasmid dilution so that the content is well mixed.
18. Put the tube with concentrated plasmid DNA back into the freezer.
19. Leave the tube with your diluted plasmid DNA on the bench and continue with the next step.

Each student should prepare one PCR reaction. Add the reagents in the order given below in a 200 µL PCR tube. Pipette very carefully, there is only a limited amount of reagent which is expensive.

PCR amplification (50 µL):

• 33 µL 1.66 x PCR mastermix (= 2x PCR mastermix with 6x loading buffer , hhis is a green solution that has both PCR master mix and loading buffer)
• 5 µL primer 1 (5 µM, check the google sheet for your PCR reaction)
• 5 µL primer 2 "
• 7 µL of the x 200 diluted plasmid DNA.

See see pTAx assembly strategy

The teacher will take the tubes to the BioRad T100 thermal cycler in the LGM laboratory.

Each group should make 100 mL liquid YPD medium in a 250 mL Schott flask. Put a small piece of autoclave tape on the lid. Label the flask properly (Content, Date, Course, Turno, Group etc.).

material#LAB3

Summary:

• Analyze PCR products from LAB2 on gel#2
• Inoculate S. cerevisiae in liquid YPD cultures with from LAB2

It is critical that you keep track of your PCR tube. They are very small and have only a number written on them. We need them for the transformation later, so we can not lose them.

1. Put the gel in the gel tray in a square petri dish. The gel should have at least one well per group number and one extra for the marker.
2. Add 5 µL of the PCR product DNA to an empty well, start with the leftmost well.
3. All group members should load their PCR product.
4. Take note of where your PCR product is.
5. The teacher will help you to put the gel in the electrophoresis chamber.
6. The teacher will then load the molecular weight marker.
7. Apply the electrical field as soon as you are done.
8. The electrophoresis last around 15 min at 200 volts in the Bachman gel tank using a homemade rectifier.
9. When the gel run is completed, the teacher with take a picture using a transilluminator.

The first task is to measure the optical density of the culture. We do this by diluting the culture ten times in the same medium and measuring the OD640 nm against the medium as blank.

1. Add some YPD medium to a plastic weighing boat. This medium does not need to be sterile.
2. Add 900 µL YPD from the weighing boat to a cuvette.
3. Mix cell culture and add 100 µL to the cuvette.
4. Measure OD640 with a spectrophotometer.
5. Calculate the optical density of the culture.
6. Calculate the volume of cells from the pre-culture that needs to be added to 40 mL of YPD medium to attain an OD640 = 0.17 (OD640 = 0.17 = 5 x 106 cells/ml using a spectrophotometer GENESYS20.)

We can calculate the culture volume we need to add by the equation below. This is a simple mass balance:

$Va \cdot OD640 = 0.17 \cdot (40 + Va)$

• Va is the volume we use to inoculate (mL).
• OD640 is the optical density for the the culture from which we inoculate.
• The culture volume before inoculating is 40 mL.
• The final optical density we want is 0.17.

If we rearrange the equation: $Va = \frac{6.80}{OD640 - 0.17}$

1. Wipe down the bench with ethanol and light the lamp to create a sterile environment.
2. Add 40 mL of YPD medium to a 50 mL FALCON tube.
3. Add $Va$ mL of pre-culture to your tube.
4. Mix and measure the OD640 by removing one mL to an empty cuvette
5. If the OD640 is ok, pour all the contents of the tube into a sterile Erlenmeyer flask.
6. Incubate until cells have grown for two generations i.e. final OD640 should be 0.17 * 2 * 2 = 0.68
7. Pour the cells into a sterile 50 mL FALCON tube and store on ice.

Each group should make 500 mL solid SD medium in a 1 L Schott flask. Put a small piece of autoclave tape on the lid. Label the flask properly, (Content, Date, Course, Turno, Group etc.).

material#LAB4

Summary:

• Wash competent yeast cells
• Preparation of DNA mixture
• Yeast transformation
• Plate transformants on solid SD medium from LAB3

42. Decant the supernatant slowly without disturbing the cells.
43. Pour the content of the yeast culture into a 1.5 mL Eppendorf tube
44. Spin for 20 s in a micro-centrifuge at top speed
45. Remove supernatant by decanting (pipette not needed).
46. Add 1 mL ultra pure water.
47. Resuspend cells by pipetting slowly with a P1000 pipette (slowly, don't make froth).
48. Spin for 20 s in a micro-centrifuge at top speed
49. Remove supernatant by decanting (pipette not needed).
50. Add 800 µL ultra pure water.
51. Resuspend cells by pipetting slowly with a P1000 pipette (slowly!).
52. Put the tube on ice.

We need two mixtures, one complete that we call "➕" and one that lacks the pBR fragment that we call "🔺" (delta)", see Table #5. The ∆ mix is a negative control where we would expect few transformants. The plan is for 12 students to transform with the "➕" mix and 4 students with the "🔺" mix.

53. Pool all successful PCR products together except the tubes with the pBR fragment.
54. Measure the volume using a pipette ( for example 380 µL)
55. Divide the mixture 3/4 (➕) and 1/4 (🔺) (285 µL and 95 µL)
56. Add the pBR fragment to the ➕ mix (70 µL)
57. Add an equal portion of water to the 🔺 mix (70/3 = 23 µL)
58. Add 500 µL water to the ➕ mix (to have enough volume for 12 x 60 µL)
59. Add 167 µL water to the 🔺 mix (to have enough volume for 4 x 60 µL)
60. Mix by vortexing.
61. Divide ➕ mix into 3 tubes, 285 µL in each (one for each group).

Each student should make one transformation. This protocol is described in detail here.

62. Mix washed cells by inverting the tube.
63. Transfer 67 µL of the cell suspension to a clean 1.5 mL Eppendorf tube per group member (If the group has four members, you need four tubes.). Mark the tubes with your initials.
64. Centrifuge the cells for 20s at the highest speed of the microcentrifuge.
65. Remove supernatant with a P200 pipette. Leave the cell pellet at the bottom of the tube, do not resuspend.
66. Add 40 µL of ➕ or 🔺 DNA mix to the tube with cells.
67. Add 200 µL PLS (PEG-LiAc-ssDNA). Be careful and pipette slowly as PLS is sticky. Use a P1000 pipette with blue tip.
68. Vortex the tubes until cells are well resuspended and no clumps visible.
69. Put the tubes in a floating tube rack at 42°C.
70. Incubate for 40 min.
71. Mark a Petri dish with the appropriate solid medium with your group number and name. Write on the back side of th eplate, not on the lid.
72. Add about 1/2 mL glass spheres (~10-15 spheres) to the Petri dish.
73. Time for ☕ break.
74. Remove tube from water bath after 40 min and put tube on ice for at least 2 min.
75. Spin tube for 20s at highest speed.
76. Remove supernatant with a P200 pipette. Leave the cell pellet at the bottom of the tube.
77. Add 300 µL YPD medium and resuspend with the pipette by slowly pipetting up and down. Be careful as the cells are sensitive
78. Transfer 100 µL of the cell suspension to your Petri dish.
79. Spread the cells by shaking the glass spheres (The samba method).
80. Give the rest of the cell suspension to the instructor
81. Incubate the plates upside down for 2-4 days at 30°C.

material#LAB5

The objective of this task is to simulate the outcome of each of the five PCR product used to make the vector.

1. The primer numbers and template plasmids used can be found in Table#4 above.
2. The template sequences are available through links in Table 1.
3. The primer sequences for each primer number can be found in Table 3.
4. Simulate the PCR products using primer sequences and
5. Add the size and ldseguid checksum for each of the PCR products to the LAB5 tab in the course Google sheet.

1. Use the Assembly simulator (here) to join the five sequences.
2. Annotate your sequence with pLannotate
3. Add the size and cdseguid checksum for your plasmid as well as the sequence to the LAB5 tab in the Course Google Sheet.

Pick one isolated colony with a sterile pipette tip and inoculate a Petri dish with the the appropriate solid medium.

material#LAB6

Summary:

• NaOH total yeast DNA preparation
• Colony PCR

Each student should have a plate with colonies. Do not contaminate this plate, we will need these yeast cells later.

1. Watch this YouTube video (2 min) describing the technique.
2. Add 20 µL 20 mM NaOH to a clean 1.5 mL Eppendorf tube.
3. Pick a small amount of cells from your plate (from LAB5) with a yellow tip. It is important not to take too much and no agarose (see at 52 s in the video).
4. Add the cells to the solution and swirl to mix (see at 59 s in the video).
5. Incubate tubes at 95°C for ten minutes.
6. When the 95°C incubation is over, add 180 µL TE buffer.
7. Vortex the tube for 30 s.
8. Spin at max speed in a micro-centrifuge for 10-20 s.

1. Add 15 µL of PCR mix^* to a new PCR tube (these are the small tubes).
2. Add 5 µL of the yeast DNA to the PCR tube, do not distrub the cell debris from the bottom of the tube.
3. Put tubes in PCR machine (or freeze the tubes at -20°C for later).
4. Run this PCR program:

Taq DNA pol
|95°C  |95°C               |     |
|______|_____          72°C|72°C |
|10min |30s  \ 53.8°C _____|_____|
|      |      \______/ 0:30|5 min|
|      |          30s      |     |

^* PCR mix for 25 reactions (15 µL for a 20 µL PCR reaction):

1. 25 * 13 = 260 µL 1.5x Green PCR mastermix
2. 25 * 1 µL = 25 µL 1222 (10 µM)
3. 25 * 1 µL = 25 µL 1779 (10 µM)

Summary:

• Run gel with colony PCR products from LAB6
• Inoculate 1 mL yeast cultures for plasmid rescue from liquid medium.
• Prepare solid LB Lennox medium for LAB8

1. Put a piece of gel in a square Petri dish
2. Load 8 µL of the the PCR product.
3. Run the gel for 15 - 20 min in TAE buffer.

While the gel is running, each group should prepare 250 mL solid LB Lennox medium in a 500 mL Schott bottle.

See if any of the previously prepared liquid YPD medium (LAB2) is still fresh? Inoculate 1 mL YPD medium in a 2 mL Eppendorf tube from the yeast plate using a sterile pipette tip.

This culture is for plasmid rescue next week.

material#LAB8 Summary:

• Prepare crude yeast DNA from cultures prepared in LAB7.
• Transform E. coli with crude yeast DNA.
• Plate E. coli on Petri dishes with solid LB medium from LAB7

1. Each student has at least one Eppendorf tube with S. cerevisiae frozen cells from LAB7.
2. Add 200 µL P1 solution to the frozen cells
3. Add 200 µl of glass beads (about one full 0.2 mL PCR tube).
4. Resuspend cells by vortexing briefly.
5. Vortex the tubes for 5 minutes using a disruptor genie.
6. Add 200 µL of P2 solution (use googles, solution has NaOH). Do this as soon as possible, the disruption of the cells liberate nucleases that can damage DNA.
7. Invert tube slowly for 3 min.
8. Add 200 µL P3 solution.
9. Invert tube slowly for 3 min.
10. Spin (centrifuge) tubes for 10 min at highest speed.
11. While the tubes are spinning, prepare one 1.5 mL Eppendorf tube with 1 mL 96-100% ethanol
12. Transfer 400-500 µL of the supernatant from the centrifugation to the tube with ethanol.
13. Invert tube slowly ~10 times to mix.
14. Spin (centrifuge) tubes for 10 min at highest speed.
15. Remove all liquid by decanting.
16. Add 1 mL 70% ethanol to the tube
17. Spin (centrifuge) tubes for 1 min at highest speed.
18. Remove all liquid by decanting.
19. Remove as much liquid as possible with a pipette, spin again if necessary.
20. Dry the DNA for 5 min at 50°C.
21. Add 50 µL of TE buffer to the precipitated DNA and vortex briefly to dissolve.
22. Label your tube with the number in the Google sheet next to your name.

1. Add the 10 µL of the plasmid DNA to the tube with competent cells, flick the tube a few times to mix. Do NOT vortex the cells at this point.
2. prepare a cup with ice/water slurry.
3. Incubate for 5-10 min on ice.
4. Heat shock in water bath at 42°C during EXACTLY 45 s ⏱️.
5. Cool the tube for 2 min in a water/ice slurry for fast heat transfer.
6. Add 300 µL pre-warmed (37°C) liquid LB medium.
7. Add 20 µL ampicillin to the cells.
8. Mix cells by pipetting slowly up and down.
9. Plate 200 µL by adding 20 - 30 sterile glass beads to an LB plate and swirl to spread the cells.
10. Incubate inverted at 37°C for 18 - 24 hours. This protocol is described in greater detail here.

• Prepare crude yeast DNA from cells with glass beads and an E. coli plasmid miniprep kif
• Transform E. coli with crude yeast DNA
• Plate E. coli on Petri dishes with solid LB-amp medium from LAB7

1. (Scrape some yeast cells off a plate)
2. (Add 1 mL water in a 2 mL Eppendorf tube)
3. (Wash cells a and add glass beads)
4. (Add 200 µl of glass beads (about one full 0.2 mL PCR tube) to a tube)
5. (Add the amount of resuspension solution to the cells indicated by the kit)
6. Vortex the tubes for 5 minutes using a disruptor genie.
7. Add the lysis buffer as soon as possible, the disruption of the cells liberate nucleases that can damage the DNA.
8. Follow the rest of the alkaline lysis mini-prep kit protocol. .
9. If the kit has an optional wash step to get rid of nucleases, use that.
10. Elute in a small volume 50 µL or 30 µL for higher concentration.

1. Add the 10 µL of the plasmid DNA to the tube with competent cells, flick the tube a few times to mix. Do NOT vortex the cells at this point.
2. Incubate for up to 30 min on ice.
3. Heat shock in water bath at 42°C during EXACTLY 45s.
4. Cool the tube for 1-2 min in a water/ice slurry for fast heat transfer.
5. Add 1 mL pre-warmed liquid LB medium to one and proceed to the next step or let cells recover at 37°C for 1 h if time allows.
6. Pipette 300 µL of the content to a new Eppendorf tube. Give the remaining cells to the teacher.
7. Add 20 µL ampicillin (x1000) and mix by inversion
8. Pipette all of the content to a LB plate with 10-20 sterile glass beads and swirl the plate to spread the liquid.
9. Incubate plates inverted for 18-24 h at 37°C

material#LAB8

Summary:

• Plasmid miniprep using kit
• Load & run agarose gel#4 on plasmid preparations.

Preparation of ~16 plasmids (One per student) using alkaline lysis mini prep. We will use the a alkaline lysis plasmid miniprep protocol. The teacher has prepared E. coli cultures in liquid or on solid medium beforehand. Cultures with each plasmid were grown in or on LB with ampicillin for selection of the plasmids. Here is a short protocol for printing.

129. Add 10 µL 6 x loading buffer to your plasmid DNA.
130. If necessary, spin the tube for ~2-3 seconds to collect the liquid at the bottom.
131. Put the gel in the gel tray.
132. Add more buffer (if needed) until the gel is just submerged.
133. Add 8 µL of the plasmid DNA to an empty well, start with the leftmost well.
134. All group members should load their plasmid.
135. Take note of where your samples are.
136. The teacher will help you to put the gel in the electrophoresis chamber.
137. The teacher will load the molecular weight marker.
138. Apply the electrical field as soon as you are done.
139. The electrophoresis last around 15 - 20 min in the Bachman gel unit powered by the homemade rectifier.
140. When the gel run is completed, the teacher with take a picture using a transilluminator.

material#LAB10

Summary:

• cut DNA with restriction enzymes
• run agarose gel#5

141. Unfreeze the plasmid solution.
142. Pipette 8 µL of the plasmid solution into a new Eppendorf tube.
143. Mark the Eppendorf tube
144. Ask your teacher to add 1 µL of Restriction enzyme buffer and 1 µL enzyme (Table#6) to your tube.
145. Spin and incubate for one hour at 37°C. Continue to assemble the plasmid in-silico.
146. Add 2 µL DNA loading buffer to your tube
147. Load all of the tube content on an agarose gel