# Fusion Proteins for Transfecting Mammalian Cells
A common technique for visualizing proteins and structures in cells is to generate artificial gene expression plasmids that contain a gene encoding a fluorescent fusion protein. In these plasmids the coding sequence for a fluorescent protein (such as Green Fluorescent Protein, GFP) is fused a sequence that encodes a protein that we want to investigate. The resulting fusion protein allows us to visualize the distribution of the protein of interest in cells.

##### Image: https://www.thermofisher.com/content/dam/LifeTech/global/life-sciences/cellanalysis/Images/0914/figure_fp_4.jpg

In the coming labs we will be transfecting the mammalian cells we have been working with (HEK293T cells) with plasmid DNA. The plasmids we will be transfecting our cells are:

### ER-RFP

The plasmid contains the sequence for a red fluorescent protein derived from a *Discosoma sp*., a genus of [disc-shapped anenomes](https://en.wikipedia.org/wiki/Discosoma), fused to the signal peptide from [prolactin](https://www.uniprot.org/uniprot/P01236) which results in the RFP protein localizing to the endoplasmic reticulum (ER).

### mEmerald-ELP-1-25

The mEmerald-ELP-1-25 plasmid contains the sequence for a green fluorescent protein derived from Aequorea victoria, the [crystal jellyfish](https://en.wikipedia.org/wiki/Aequorea_victoria), fused to the [human ER lumen protein-retaining receptor 2, ELP1](https://www.uniprot.org/uniprot/P33947) which results in the fusion protein localizing to the golgi.

## Objectives of the lab:
This week you will be isolating plasmid DNA from bacteria for transfecting your cells. Upon completion of this lab, you should be able to:
1) Isolate plasmid DNA from bacterial cells
2) Load and run a gel to visualize your plasmid DNA.
3) Perform a restriction digest to confirm the presence of your fusion protein and determine the concentration of your DNA.

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## “MiniPrep” Plasmid DNA purification
### How it works
There are many methods to isolate plasmids from bacteria, and we will use one of the most common one. This method relies on alkaline lysis of the bacterial cells. A popular kit for plasmid isolation, the NucleoSpin Plasmid DNA Purification Miniprep kit will allow us to purify the plasmids after lysis by using the silica columns, with which you are now familiar. Bacterial cells are centrifuged into a pellet and resuspended in a small volume of saline buffer. Then they are lysed (ruptured) by the addition of a strong ionic detergent, sodium dodecyl sulfate (SDS), and a strong base, sodium hydroxide (NaOH). The cell membranes will rupture by being solubilized by the detergent SDS, while the strong base denatures DNA and proteins. RNase A, an enzyme that digests RNA, is also included in the mix to remove RNA. All this happens in less than 5 minutes! The lysate is neutralized by the addition of an acidic, high salt solution (guanidine hydrochloride), which also causes precipitation of proteins, membrane fragments, SDS, and the bacterial chromosomal DNA. When this residue is centrifuged, the remaining supernatant should contain re-natured plasmid DNA, small soluble molecules, and some residual protein. Next, the plasmid is isolated using a silica-resin DNA-binding column. Once the plasmid DNA is bound to the silica matrix it will be washed with solutions containing ethanol and then finally eluted in an aqueous solution. This method produces microgram quantities of plasmid DNA, so they are known as “minipreps”.

As the bacteria replicate the circular, double stranded plasmid DNA, the plasmid is enzymatically twisted upon itself. We call this supercoiled DNA. This coiling ‘packages’ the plasmid DNA into a physically small shape. However, during plasmid isolation (the miniprep) a small amount of the plasmid DNA may be mechanically damaged by pipetting. This mechanical damage will result in some plasmid molecules that receive either a double-stranded break, forming a linearized plasmid (linear form), or a single-stranded break, forming a ‘nicked’ circular plasmid without any supercoils (nicked form).

Additionally, if the DNA spends more than a few minutes in the strong alkaline solution, some plasmids may not re-nature, producing a single strand form of the plasmid. Therefore, the recombinant plasmid DNA isolated from the transformed DH5 cells in your overnight culture will contain a large concentration of only one plasmid sequence, but plasmids will be in different structural forms. These forms of the plasmid will actually provide useful information, and they need to be considered when you analyze your undigested recombinant plasmids using gel electrophoresis.
## Restriction Enzyme (RE) Digestion
Restriction endonucleases are commonly used in molecular techniques to confirm the identity of DNA samples, for genetic mapping, and as one possible way to clone pieces of DNA into plasmids. Restriction enzymes will only cut double-stranded DNA, by binding to a unique DNA recognition sequence and breaking (“cutting”) the covalent bond between two nucleotides at a specific position on both DNA strands. Recognition sequences, on average, range from 4 bp in length up to 8 bp or more in length, with a 6 bp sequence being the most commonly used in cloning techniques due to the frequency with which the recognition sequence is found in DNA. While using restriction enzymes to verify a DNA sequence is not as rigorous as DNA sequencing, it is much faster and less expensive.
## Agarose gel electrophoresis
Up to this point we have assumed that the genomic DNA isolation, PCR and purification have worked as expected. While these procedures usually work well, unexpected results can occur, and it is always good laboratory practice to verify the outcome of procedures before investing more effort in samples. Moreover, it is important to estimate the concentration of your purified PCR product in order to use an appropriate amount in the recombination reaction next week. Electrophoresis will separate DNA fragments by their size and allow a relative estimate of their concentration.
### How it works
DNA is negatively charged, due to the phosphate groups in its backbone. This charge allows electrical current to move DNA. Agarose is a polysaccharide polymer that forms a porous gel matrix. Differently sized DNA fragments can be separated by placing a current across the agarose gel. Negatively charged DNA will migrate toward the positive electrode. (Electricians mark positive electrodes with red, so the DNA will “run to the red.”) Smaller DNA fragments will fit through the porous gel more easily, moving farther through the gel than larger fragments in the same amount of time. Large fragments cannot fit through the gel matrix as easily or as quickly. The DNA in the gel is revealed by a fluorescent dye that binds to nucleic acids included in the gel. The natural fluorescence of these dyes increases 50-fold when bound to DNA. Some common dyes used to stain for DNA are ethidium bromide (EtBr), GelStar (from Lonza), and SYBR Green (from Life Technologies).
If DNA samples are loaded into a well at the top of the gel, electrophoresis moves DNAs of different sizes different distances, producing distinct bands of DNA in the gel. Each band consists of many DNA molecules of the same size -- the same number of nucleotides.
Several conditions can affect how your DNA travels through the gel. Depending on the expected sizes and number of bands that altering these conditions may improve the resolution of the gel.
* Agarose Concentration (0.5-2% is the normal range used)
* Voltage (50-275V depending on the type of gel buffer used)
* Time (6 minutes to ~1.5 hours, dependent on voltage and the length of the gel)
* Amount of DNA loaded in a lane
## Estimating size and mass of DNA
The approximate size of the DNA in a band (also called its length) is determined by comparing the position of the band in the gel relative to other bands with known sizes. A lane containing size standards called a DNA ladder is always included among the samples run on a gel. Different DNA ladders are available with bands in different size ranges.

DNA does not move through the gel in a linear relationship to its size. The distance DNA moves is related to its size logarithmically. Be aware of this fact when you estimate sizes. Simply put, smaller DNAs run a lot farther and faster than large DNAs. While it's not possible to estimate exact sizes (e.g. 1745 bp), your estimate should have two significant digits (e.g. 1700 bp).

The mass of DNA in a band can also be estimated from the gel. Compare the intensity of bands in the DNA ladder to your sample bands. The fragments in the DNA ladder have known masses, as well as known sizes. The more DNA is in a band, the more fluorescent dye (like GelStar) is bound and the greater the fluorescent intensity of that band. One caveat is that a single GelStar molecule will bind about every 50 bp. Larger DNA fragments bind more GelStar molecules, making them brighter than smaller fragments even if the same mass is contained in each band (For example in Figure 1, compare the 10 kb and 0.5 kb bands.) Therefore, the best estimate of mass in a sample comes from comparing your sample band to the brightness of a ladder band of similar size.

From the mass of the DNA found in the band, it is possible to estimate the approximate concentration of that DNA in your original sample. The concentration of DNA in your original sample (ng/μl) can be found by dividing the estimated mass of DNA (ng) by the volume (μl) of the original sample loaded on the gel. For example, imagine you used 3 μl of PCR reaction to prepare a 5-μl sample to load on your gel. After running the gel, you estimated that there was ~200 ng of a ~700 bp PCR product, suggesting a concentration of ~70 ng/μl in the original PCR sample (200 ng PCR product / 3 μl of PCR product). Keeping the number of significant figures (1) rounds the answer to 70 ng/μl.
## The Lonza FlashgelTM system
In order to save a little time, we will use pre-made agarose gels that are designed to run at high voltages for shorter times. This gel system is made in Rockland, Maine by Lonza and utilizes a FlashGel dock containing a light that can illuminate DNA bands (bound to the GelStar fluorescent dye) without the need for a separate UV light source. This will allow you to watch the DNA in your samples separate in real time. The 1.2% agarose gel cassettes contain 13 wells and can withstand the higher temperatures produced when running a gel at higher voltages (up to ~275V). Because samples will be run quickly (6-7 minutes), we will be unable to separate DNA fragments larger than ~3 kb. Since all of our PCR products are expected to be smaller than 3 kb, this should not be a major issue. Other drawbacks of using this system include a higher cost per gel and a shelf life of ≤ 6 months for the gel cassettes.

We will use two DNA ladders. The 1-kb ladder from New England Biolabs (Figure 1) enables estimates of mass for your samples. The 100-4000 bp DNA ladder from Lonza (Figure 2) contains 4 additional reference bands in the 100-1000 bp range to provide more accurate size information for smaller DNAs. However, this ladder lacks mass information, making it uninformative for estimating DNA concentrations.

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### A note on concentrations
Many commonly used lab solutions, like buffers and loading dyes, contain multiple compounds at different molar concentrations. Therefore, rather than listing the concentrations of each ingredient, researchers typically define the working concentration of the solution as “1X”. So, preparing large amounts of these reagents, it is often convenient to make more concentrated stock solutions. A 5-fold more concentrated version of loading dye would be labeled “5X”. It can then be diluted 1:5 to provide a 1X working concentration in your sample. Keep an eye out for this notation, and be careful not to use a stock when you need a 1X working concentration!

# Procedures
## Minipreps
Each research team will receive two overnight bacterial cultures of DH5α E. coli that contain the plasmids for this lab. Apply this procedure to both of your overnight cultures.

***Note: Many of the solutions in this procedure are irritating to the skin, so wear gloves.***

1. Balance the swinging bucket centrifuge and spin your overnight cultures at 3000 rpm for 5 minutes (without the snap cap lids).
2. Pour the supernatant into a liquid waste container, containing bleach. Allow it to drip for a few seconds onto paper towels for drainage. There should not be more than a damp residue of broth on the pellet.
3. Resuspend the bacterial pellet in 250 μl of P1 buffer by gently pipetting up and down. Then, using the same pipette tip, transfer the suspension to a clean 1.5 ml microcentrifuge tube.
4. Add 250 μl of P2 buffer to the bacterial suspension. Invert the tube several times to mix.
* During this step the bacteria are being lysed. The samples must be mixed carefully at this point to avoid shearing the bacterial chromosomal DNA.
***Quickly move to the next step.***
6. Add 350 μl of N3 buffer and immediately invert to mix several times.
* This buffer contains guanidine hydrochloride, a chaotropic denaturing agent. You should notice proteins precipitating out of solution. Adding this solution quickly to neutralize pH, minimizes the formation of single-stranded plasmid.
7. Centrifuge your sample at maximum speed (≥14000×g) for 10 minutes.
***Carefully remove the tube from the centrifuge.*
Dislodging the pellet may add contaminants to your DNA in the supernatant.**
8. Carefully remove all of the supernatant using a pipette. Transfer it to a spin column.
9. Centrifuge the column at maximum speed for 1 minute.
10. Discard the flow-through into a small waste container. Do not combine with bleach waste!
* The flow-through contains guanidine hydrochloride, which will react with bleach (sodium hypochlorite) to form poisonous chlorine gas.
11. Wash the column with 750 μl of PE buffer.
12. Centrifuge your sample at maximum speed for 1 minute.
13. Discard the flow-through into a small waste container
14. Centrifuge again at maximum speed for 1 minute. ***This is critical to remove residual ethanol from the column.***
15. Transfer the column to a clean, labeled 1.5 ml microfuge tube
16. Add 50 μl dH2O to the center of the column.
17. Incubate for 1 minute at room temperature.
18. Centrifuge your sample at maximum speed for 1 minute. It is okay that the lids on your microcentrifuge tube are not capped, just position them in the rotor so that the lids do not snap off from the force of the spin.

***You have now eluted your plasmid DNA. Some of this sample will be used for the restriction enzyme digestions and gel electrophoresis performed next. The remainder should be stored in the freezer for transfection into HEK293T cells next week.***

## Restriction endonuclease reaction
You will set up one double-digest (a DNA digest with two restriction endonucleases) on each of your miniprep DNA samples. The digest will be with the restriction enzymes XbaI and BamHI. Using the restriction enzyme maps of you plasmids, estimate the expected number of bands and band sizes that will result from your plasmid digestion for each plasmid:

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#### ER-RFP

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**Restriction enzymes used for digest: ______________________
Expected # of bands: ______________________
Expected band sizes: ______________________**

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#### mEmerald-ELP-1-25

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**Restriction enzymes used for digest: ______________________
Expected # of bands: ______________________
Expected band sizes: ______________________**

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1. For each digest, mix the reagents listed in the recipe below for each sample.
2 µl mini prep DNA
16 µl dH2O
2 µl CutSmart Buffer
20 µl total volume
2. Add 1 µL of the BamH1 restriction enzyme.
3. Add 1 µL of the XbaI restriction enzyme.
4. Mix by pipetting up and down. If needed, spin the contents of the tube down.
5. Incubate for ~15 minutes at the appropriate temperature for the enzyme.

## Sample preparation for the Lonza FlashGel
In new microcentrifuge tubes, prepare each of the following samples for the gel according to Table 1.

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1. Prepare each of your samples for the gel, according to Table 2.
• ***DO NOT*** add loading dye to your original samples! It will prevent future steps (like transfection of mammalian cells) from occurring.
• Check to be sure each sample is as ‘purple’ as the DNA ladders to be sure enough loading dye was added. If the sample is a much lighter purple, add another microliter of 5x loading dye.

*In order to monitor the progress of the electrophoresis, 5X loading dye (blue color) must be added to your samples. These visible dyes are negatively charged, and their movement through the gel reflects the movement of DNA. The dye solution also contains a dense, chemically inert compound (such as glycerol or sucrose) to allow the samples to sink to the bottom of the well during loading.*

2. Line your samples up in the rack in the order you will load them on the gel.
*Record this order in your notebook.*

*Once the loading dye has been added to a sample, it is not necessary to keep it on ice any longer. A good practice is to line up all of your gel samples in the order that you want to load them before you start so that you can quickly load your gel. You want to be as timely and efficient as possible so that the samples do not diffuse into the buffer.*

3. Freeze your original, undigested miniprep DNA samples for the next lab.
* Each tube should be labelled with the name of the plasmid, the date and the initials of each member of the reseach team.

## Loading and running the Lonza FlashGel
**Read through all of these steps first!**

The entire lab will share a gel and should load the gel at approximately the same time. The DNA in samples will begin to diffuse if left too long in a well with no current applied. When you load the gel, leave at least one well on both the right and left sides without sample as the current will sometimes create gel artifacts in the outermost lanes.

### To prepare the FlashGel
1. Open the FlashGel package and remove the white sticker from the top of the cassette.
2. Over the sink, flood the wells gently with dH2O and tilt the cassette to pour off any excess water. (It is important to leave liquid in every well.)
3. Insert the cassette into the FlashGel Dock.
4. Load your samples. Use the tips below.
* If you have fewer samples than number of wells (13 total in the Lonza flashgels), avoid loading the wells on either end of the gel.
5. Place the pipette tip at the top of the well, below the level of the fluid.
* Expel your sample slowly.
* Move the pipette away from the well, before you release your thumb. (Otherwise, you risk pulling your sample back up into your pipette tip.)

**Try to have all samples loaded within ~1-2 minutes.**

6. Connect the leads to the power supply. Then, turn on the power.
7. Set the voltage to ~150V (with the range select set to high).
* Your instructor may ask you to change the voltage depending on the time available, to allow for better separations and more accurate mass estimations. At ~150V, it should take 15-20 minutes for your samples to run through the gel. At ~100V (with range select set to low) it will take about 30 minutes but you will getter better resolution of larger fragments.
8. Periodically, turn on the dock light to watch your DNA travel through the gel. You may wish to capture an image about halfway through the electrophoresis while the gel is still running (see step 9).
9. When the DNA ladders are well separated and the smallest band (ladder band or smallest digested fragment) is near the bottom of the gel, turn off the power supply and disconnect the leads.
10. Take a picture of your gel using your cell phone camera.

## Gel Analysis
When interpreting your gel results, there are several facts to keep in mind. First, remember that the relationship between a DNA fragment’s size and its migration distance is logarithmic—not linear. So instead of evenly distributed bands in the DNA ladder, smaller fragments separate more (and their bands are more spread out), while larger fragments separate less (see Figure 1). This means that the resolution of an agarose gel is better for smaller DNA fragments than for the larger ones. Therefore, your size estimates will be more accurate for smaller fragments.

Also be aware that if a well is loaded with too much DNA, bands will be more spread out (wider), as lots of co-migrating DNA pushes against its neighboring molecules. The accuracy with which you can estimate the size of these heavy bands is limited, although the upper edge is often the best indication of the DNA's true size.

Finally, when you have two different DNA fragments that are close in size, if they are not separated effectively they may appear to be a single fat band instead of two thin bands. Taking more than one photograph of your gel at different exposures can sometimes help distinguish among similar bands. While there are many variables with gels, if you take your time analyzing the results, a lot of useful information can be gained.

1. Obtain your gel image and import it into your lab notebook for labeling.
2. Above the gel image, label each lane so it is clear which sample you are looking at easily.
3. Label the size of each band in the DNA ladders.
4. Label the correct mass of each band in the DNA ladder.
*Remember that the mass for the DNA ladder bands is for a 10 μl volume loaded in the gel, while you only loaded 4 μl of the ladder in your gel. Be sure to compensate for this volume difference.*
5. Estimate the size of the band(s) in your samples, by comparisons to the DNA ladders (Figures 1). *Interpret the results that you actually observe—not the results that you expect to see.*
6. Compare the observed results to the expected results.
* ***What size bands are you expecting from your digested plasmid samples?***

7. Estimate the concentration (in ng/μl) of any bands in your digested plasmid samples by comparing the intensity of the products to the intensity of the 1 kb DNA ladder (Figure 1). Keep in mind that you loaded 4 ul of the original digest (excluding the volume of loading dye because it contains no DNA).
8. Estimate the concentration of your original miniprep samples.
* For each of your digests, you added 2 ul of your original mini prep plasmid sample in a total of about 20 ul in each digestion.
* ***What is the dilution factor that you need to multiply by in order to obtain the concentration of your original mini prep sample?***

* A dilution factor is the total volume of a sample plus diluent after dilution divided by the initial volume of sample.
Ex: 100 mL of final volume ÷ 2 mL original volume of sample = 50 dilution factor*