# An Introduction to Mammalian Cell Culture
Mammalian cell culture is the process of growing mammalian cells that have been isolated from an organism in flasks or dishes in the lab. This technique is widely used in biology labs as it allows researchers to easily manipulate cells under controlled conditions. Cultured cells have been used extensively to study a wide variety of cellular processes with higher resolution than is possible in intact organisms. Cultured mammalian cells are also used to produce vaccines and antibodies for therapeutic uses.
## NIH3T3 Cells
The cells we will be working with are NIH-3T3 cells. NIH-3T3 cells are a fibroblast cell line that was isolated from a mouse NIH/Swiss embryo. The cell line was initially established in 1962 at New York University. NIH-3T3 cells are relatively fast growing cells that are fairly easy to genetically manipulate. These qualities have made NIH-3T3 cells extremely useful and, as a result, the use of NIH-3T3 cells is widespread and the cells are commonly found in research labs today.

<a title="Gleiberg, CC BY-SA 2.0 DE &lt;https://creativecommons.org/licenses/by-sa/2.0/de/deed.en&gt;, via Wikimedia Commons" href="https://commons.wikimedia.org/wiki/File:NIH3T3-Zellen.jpg"><img width="256" alt="NIH3T3-Zellen" src="https://upload.wikimedia.org/wikipedia/commons/thumb/2/25/NIH3T3-Zellen.jpg/256px-NIH3T3-Zellen.jpg"></a>
"File:NIH3T3-Zellen.jpg" by Gleiberg is licensed under CC BY-SA 2.0.

## Objectives of the lab:

This week’s lab is designed to introduce you to mammalian cell culture. Upon completion of this lab, you should be able to:
1) Use a light microscope, serological pipets and micropipettes
2) Passage/manipulate mammalian cells
3) Count cells using a cell counting chamber (hemocytometer)
4) Calculate cell densities
5) Develop and test a hypothesis as to what a given serum concentration does to a human cell
6) Create a graph of your data with error bars

## Part 1: Microscopy, Cell counts and Diluting cells

### Parts of a Microscope


*Figure 1. Zeiss Axio Vert.A1 FL Microscope*

**Ocular:** The piece you look through. Sometimes called an ocular lens or eyepiece, this unit is really a series of lenses. Our microscopes are binocular, having two oculars. Learn to use both eyes; focus your eyes as if you were looking at an object about five to ten meters in front of you. You should adjust the width of the oculars to match the width of your eyes.

**Diopter:** When you look through a microscope with two eyepiece lenses, you must be able to change the focus on one eyepiece to compensate for the difference in vision between your two eyes. The diopter adjustment does this. The way to correctly adjust the diopter on a lens is to first close the eye over the eyepiece with the diopter adjustment and normally focus the microscope so that the open eye sees the image in focus. Next, switch eyes (close the open eye, open the closed eye) and without changing the main focus knobs, focus on the image by turning the diopter adjustment only. Now with both eyes open, the image should be clear with both eyes. (This technique is used with binoculars too.)

**Objective lens:** Sometimes called the objective; a set of self-contained lenses. The objective gathers light from the specimen and directs it through the tube to the oculars. These scopes have three objectives (4X, 10X, and 40X).

**Nosepiece**: The rotating turret to which **objectives** are mounted. There are preset positions for each objective, detected by slight pressure changes while turning the nosepiece and usually a clicking noise. You should not grab the objectives to turn the nosepiece - use the grey ring instead.

**Stage:** The flat surface upon which slides are placed. On your microscopes, the stage moves up and down and the slide is manipulated by a geared device. A moveable stage is sometimes called a mechanical stage. The slide is moved left/right and front/back by two knobs projecting downward from the stage.

**Condenser:** A lens system under the stage that gathers light from the light source and focuses it on the specimen. There is a **diaphragm** in one part of the condenser that can be adjusted to allow the viewer to see different parts of the cell when using bright field illumination.

**Adjustment (Focus) Knobs:** Both coarse (large) and fine (small, inner) adjustment knobs are found on both sides of our microscopes. Remember that the coarse adjustment is used only with the low-power objective. These control a gear mechanism that raises and lowers the stage.

Other parts that found on microscopes include: the arm, the base and slide holders.

### Different Types of Microscopy: Bright Field, Dark Field, and Phase-Contrast

The type of illumination that we will be using with our microscopes is called bright field. Think of the light source as producing a solid tube of light that travels up to and through the condenser. When you view specimens with all of this light, you are using bright field illumination. Dark Field (not used in today’s lab with our microscopes): Dark field illumination seems like an oxymoron, but in this case it describes an unusual way of viewing specimens in some compound microscopes. The light that passes directly through the condenser does not enter the objective lens. Only light that has been scattered or reflected by the specimen enters the objective. As a result, you wind up seeing bright objects on a dark background. Phase-Contrast (not used in today’s lab with our microscopes): Phase-contrast microscopy allows us to see otherwise transparent organelles and structures. In a phase-contrast scope, the light hits the specimen and some of the light continues in a direct path. Other portions of the light pass through membranes which redirect the light. This redirected light is slowed down by 1/4 a wavelength (a phase shift of 1/4) by passing through a special filter. This special filter is shaped like a doughnut and is called a phase ring. The redirected and out of phase light eventually reaches your eyes but not at the same time as the unaltered light that passed straight through. The end result is that you can see transparent structures because they altered the pathway of light as it went through the structures. This allows us to view subcellular structures within living cells.

### Use of the Zeiss Axio Vert.A1 Fluorescence Microscope
*Note: this is an inverted microscope, so samples must be positioned on the stage so that they are facing the objectives.*
1. Position your chair so that the scope so it is directly in front of you and your chair is adjusted so that you do not have to strain to view a specimen.
2. Power the microscope on with the black power switch located on the left hand side of the base of the microscope.
3. Ensure that the filter wheel is set with the white line on Brightfield (BF).
4. Adjust the diaphragm (red arrow) for proper amount of light to visualize your sample.
* Furthest to the left = most open; furthest to the right = closed


5. Ensure that the microscope is set with the 10X objective in the "in use" position.
6. Place your sample on the stage
* If you are viewing a sample in a dish, the dish may be placed carefully on the stage over the objectives.
* If you are viewing a sample on a slide, the slide should be placed carefully on the stage over the objectives with the coverslip facing downward toward the objectives

7. If necessary, adjust the position of your sample so that it is over the objective using the XY stage controls on the right side of the microscope.


8. Use the coarse adjustment to lower the stage slowly while looking through the oculars until the specimen comes into focus. Adjust the focus to its sharpest with the fine adjustment
9. Readjust the light intensity to reduce glare and center the specimen in the field of view by moving the stage.
10. Adjust the condenser's diaphragm to maximize the resolution of the structure you are trying to see. The actual setting will depend on what you are trying to see. Small translucent objects will be seen more easily with the diaphragm closed substantially while large pigmented structures are easier to see with the diaphragm wide open.

---

**OBSERVATION In your notebooks, draw your cells as seen through the microscope**

---
# Counting Mammalian Cells
## To Trypsinize and count viable cells with a hemacytometer:

*Figure 2. Cell concentrations (cells/mL) quantified with trypan blue exclusion using a hemacytometer. A) A hemacytometer has two counting chambers per slide with a special coverslip that allows 10 µL sample to spread out underneath the coverslip in a single layer of cells. B) Etched onto the metal plate is a grid, viewed here with a 4x objective. Each counting area is 1mm2 with the central counting area shown in blue and the four corners shown in red. Note that each counting area itself is broken up into a smaller grid which can vary from the central 5x5 grid to the corner 4x4 grids. Using a 10x or 20x objective will allow you to see a single entire counting area at a single time which makes it easier to count. C) Viable cells are unstained and have a slight ‘halo’ around them due to the refraction of the intact cell membrane. Nonviable, or dead cells, have absorbed the trypan blue dye and appear blue in color and lack the ‘halo’. Cell size and shape can vary within cultured mammalian cells though some may have a round appearance.*

1) Obtain a 35mm dish containing 3T3 cells.
3) Remove the culture media from your 35mm dish using a p1000 Pipette with a clear tip (liquid and solid waste containers are provided at each bench for proper disposal).
4) With a new pipette tip, add 500 µL of Trypsin solution and gently rock the plate so that all cells are covered.
* *Trypsin cleaves proteins attaching cells to the plate*
6) Incubate the plate for at 2-3 min with occasional swirling. Cells will visibly start to release from the plate so look for the cell monolayer to lift and break up before moving to the next step.
7) Add 500 µL of 1xDMEM cell culture media to the dish and mix your cells by using the same clear tip to pipette the solution up and down around the dish. This will ensure that all of your cells have lifted from the bottom of your dish and that clumps of cells are broken apart in the solution for counting.
8) The final volume of cell suspension is approximately 1 mL. Transfer 500 µL to a new microcentrifuge tube.
9) In a second microcentrifuge tube, add 100 uL of 0.4% trypan blue and then add 100 uL of your cell suspension. Gently mix the solution by gently pipetting up and down with the pipette type.
* *Trypan blue is a blue dye that can only pass through the membranes of dead cells. Therefore, the viable (alive) cells, clear or unstained, are counted because the dye was excluded.*
10) Load 20 µl of cell/dye mixture into a single chamber on the hemacytometer slide with the coverslip already in place (Fig. 2A). Your partner’s sample can be loaded into the other chamber. The blue cell solution will wick under the coverslip and cover the counting grid area. Be extremely careful of the coverslips with the hemacytometers! They are a special thickness and very expensive to replace if broken or lost.
11) Place the hemacytometer on the stage of the microscope and focus on the gridded area, first with a 4x objective and then with either a 10x or 20x objective. Either a 10x or 20x objective should allow you to see a single counting area (Fig. 2B, blue or red area) in its entirety.
12) Using a cell counter, count the total number of viable cells (Fig. 2C) in the 5x5 grid in the center of the hemacytometer grid (Fig. 2B, blue area). Reset the counter to zero, and count the four 4x4 grids in the corners (Fig. 2B, red areas) for a total of five cell counts per sample.
13) To clean the hemacytometer, carefully rinse the slide and coverslip under running water in the sink. Spray both well with 70% ethanol and gently pat dry with paper towels. Place back in the hemacytometer case for the next group. Be extremely careful with the coverslip to avoid damaging it! Yes, this should sound like a broken record!
14) Calculate concentration of cells in solution (cells/mL) by averaging the counts from your five grid areas and multiply by 10^4 and multiplying by your dillution factor. (In this case you combined your cells 1:1 with Trypan blue, for a dilution factor is 2.)

**Sample problem**
If the cells counts from a 35mm dish were 36, 16, 36, 44, and 23 in the five grid areas:
Average number of cells = ___________
Concentration of cells (cells/mL) = ___________

---

**Record your data in your notebook:**


##### *Figure 3. Calculating dilutions uses a simple algebraic calculation. In the above example, the only value not known is the volume of the stock (10 cells/mL) needed to make a 2 mL volume of a 4 cell/mL sample. This equation becomes 10x = 24. Therefore x = (24)/10 = 0.8 mL (since V2 was in mL, V1 will be in mL). Since there are 1000 µl in 1 mL, x = 800 µl. To make this diluted sample, 800 µl of the original stock would be added to 1.2 mL (1200 µl) of media to give a total volume of 2 mL with a final concentration of 4 cells/mL.*
---
## Part 2: Experiment: Serum and Cell growth
Several factors can impact cell growth rates including temperature, pH, salt, drug exposure and nutrient availability. In today’s lab you will be testing the effects of nutrient availability on cell growth rates. You will work in pairs as part of a research team. Each table of students in the lab will act as a research team consisting of 2 students. Each team will be provided with cell media containing 3 different concentrations of fetal bovine serum (FBS): 0%,2% & 5% as well as control media (10% FBS) and 2 T25 flasks of NIH-3T3 cells. Each team will compare the different media combinations with the control media. The control serves as a positive control for cell growth in your experiments.

As a research team, formulate a hypothesis about the effects of serum concentration on the rate of cell growth over time. Record your hypothesis AND draw a graph of what you expect your data to look like in your notebooks.


*Figure 4. Experimental set up to test the effect of serum concentration on the growth of NIH-3T3 cells. Cells should be seeded in each well at 0.2x10^5 cells/mL in 500 µL volume in either the control media or the media with reduced serum concentrations. Two wells of cells will be trypsizined and then combined to count each sample (e.g. two blue wells will be combined to count and then two purple wells will be combined for a count). The group will count each time point in duplicate (e.g. column 1A and column 1B for the first time point). Repeat for the second and third time points.*

For this growth curve experiment, each group of 2 students on the research team will need to trypsinize and count the cells in their T25 flask and then dilute those cells to generate two samples to seed cells in a 24 well plate (Fig. 4):

* **CONTROL:** total volume of 13 mL of cells with a concentration of 0.2x10^5 cells/mL in CONTROL MEDIA.
* **EXPERIMENTAL:** total volume of 13 mL of cells with a concentration of 0.2x10^5 cells/mL in X% FBS/MEDIA (either 0%, 2% or 5% FBS).

1) Trypsinize and count the cells in a T25 flask in a cell culture hood

* As the cells will be plated into a 24-well dish and grown for several days, it will be important to work with the cells in the flask using sterile techniques. A hood with a lot of air flow will be used to help prevent contamination of your cells and you should practice sterile techniques when working with your cell culture.
* Turn the blower on in the cell culture hood (and the light if needed). Wipe down the counter surface of the hood with 70% ethanol.
2) Carefully pour the culture media from your T25 flask into the liquid waste container containing bleach. Note which side of the flask the cells are adhered to before proceeding!
3) With a 1 mL sterile serological pipette, add 1 mL of Trypsin and gently rock the plate so that all cells are covered by a thin layer of the solution.
4) Incubate the flask flat for 2-3 min with occasional swirling. Cells will visibly start to release from the flask so look for the cell monolayer to lift and break up before moving to the next step. *Note, you can hold the flask and gently tap the side with the palm of your hand to encourage the cells to release from the plastic.
5) Add 3.5 mL of 1xDMEM media to the flask with a 5 mL sterile pipette. With the same 5 mL pipette, mix your cell suspension by gently pipetting some of the solution up and back down the side of the flask where the cells were adhered. This will ensure that all of your cells have lifted from the bottom of your flask and that clumps of cells are broken apart in the solution for counting.
6) The final volume of cell suspension should be 4.5 mL. Using a 1 mL serological pipette, transfer between 200-500 µl of the cell suspension to a new microcentrifuge tube in the hood.
7) You can count the cells on the bench top. So, in a second microcentrifuge tube, add 100 µL of 0.4% trypan blue and then add 100 µL of your cell suspension that you just transferred to a microcentrifuge tube. Gently mix the solution by gently pipetting up and down with the pipette type.
8) Load 20 µl of cell/dye mixture into a single chamber on the hemacytometer slide with the coverslip already in place (Fig. 2A). Be sure the hemacytometer is cleaned well first. If not, rinse with 70% ethanol, pat dry carefully, and then let it air dry. Be extremely careful of the coverslips with the hemacytometers!
9) Place the hemacytometer on the stage of the microscope and focus on the gridded area with either a 10x or 20x objective.
10) Using a cell counter, count the total number of viable cells (Fig. 2C) in the 5x5 grid in the center of the hemacytometer grid (Fig. 2B, blue area). Reset the counter to zero, and count the four 4x4 grids in the corners (Fig. 2B, red areas) for a total of five cell counts per sample.
11) Calculate concentration of cells in solution (cells/mL) by averaging the counts from your five grid areas and multiply by 10^4. (In this case you combined your cells 1:1 with Trypan blue, for a dilution factor is 2.)

**Dilluting your cells:**
For your experiment today, you will need to dilute your cells to a final concentration of 0.2x10^5 cells/mL for both the control samples and the experimental samples. Use the M1V1=M2V2 equation (Fig. 3).

1) Calculate the volume of trypsinized cells you need to generate 13 mL of cells with a concentration of 0.2x10^5 cells/mL using the cell concentration from your T25 determined above.

*Example: If you start with a cell concentration of .5x10^5 cells/mL then:*

*.5x10^5 cells/mL X mL = 0.2x10^5 cells/mL 13 mL
X = 5.2mL of cells*

2) Calculate the volume of media you need to add to generate 13mL of cells with a concentration of 0.2x10^5 cells/mL using the following equation:

**Total Volume you want (mL) - Volume of Cells you need (mL) to add
= Volume of media you need (mL)**

*Example: If you want 13 mL of cells then:*

*13 mL – 5.2 mL of cells = 7.8 mL of media*

Calculate the Volume of cells and media you will need to generate a total volume of 13 mL of cells with a final concentration of 0.2x10^5 cells/mL based on your previous cell counts. Record your data and check your answer with your instructor before proceeding:

3) Label a 15 mL conical tube for your control cell dilution. In the flow hood, add the appropriate volume of cells from your T25 cell suspension using a 5 ml serological pipette. Next add the volume of control media calculated above to obtain a final volume of 13 mL.

4) Label a 15 mL conical tube for your experimental cell dilution. In the flow hood, add the appropriate volume of cells from your T25 cell suspension using a 5 ml serological pipette. Next add the volume of reduced serum media (experimental) calculated above to obtain a final volume of 13 mL.

5) Obtain and label a 24 well plate with: today’s date, your initials and your Team name as well as which wells are controls and which are experimental (include the % FBS being tested).

6) Aliqout 500 µL of your diluted CONTROL cells/media into each of the wells in rows A and B of the 24 well cell culture plate provided using a 5 or 10 mL pipette (Fig. 4).

7) Aliqout 500 µL of your diluted Experimental cells/media into each of the wells in rows C and D of the 24 well cell culture plate using a 5 or 10 mL pipette (Fig. 4).

8) Be sure your lid is on the plate properly and carefully place your plate in the 37°C CO2 incubator in the cell culture room. An atmosphere of 5% CO2 is used to maintain a neutral pH in the media indicated by the red color of the media. As cells grow and respire, the media will become more acidic, turning orange and then yellow over time.

9) Your cell counts for time 0 are now 0.2x10^5 cells/mL in all samples. To generate a cell growth curve, each group will need to come back and trypsinize and count their cells 3 more times. Each group will determine when they will come into count the cells. The specific time you come in to do the counts is not important, just record the times.

* Timepoint 1: Tomorrow evening
* Timepoint 2: The next morning (or evening)
* Timepoint 3: The next morning (or evening)

10) Take your plate out of the incubator and view the cell monolayer briefly for a control well and an experimental well using the inverted scope and the 20x objective. Note any qualitative differences in the cells and media.

11) To trypsinize and count your samples at each time point, add 200 µL of trypsin (use a p1000 pipette and clear tips) to each of the wells for that time point. This should include 4 wells in the control sample (Fig. 4, blue and yellow wells for example) and 4 wells in the experimental sample (Fig. 4, purple and orange wells for example). Replace the lid.

12) Wait 2-3 min for the cells to lift from the bottom of the plate. Add 300 µl of media to each of the wells containing trypsin (use a p1000 and clear tips).

13) Using the p1000 pipette with the clear tips, pipette the cells up and down to mix and combine two wells (Fig.4, the two blue wells for example). You should have a volume of about 1 mL now combined. Transfer all of the solution to a microcentrifuge tube which has the top labelled to indicate what sample is contained, e.g. Control 1A.

14) Repeat for all 4 samples (Control 1A, Experimental 1A, Control 1B, Experimental 1B) for that time point. Use a new pipette tip for different samples!

15) Replace the lid on your 24 well plate and place back in the incubator until the next time point.

16) On the benchtop, count the cells as before. Add 100 µL of cells to 100 µL of trypan blue and pipette up and down to mix (use a p1000 pipette and clear tips). Add 20 µL to a chamber on the hemacytometer and count 5 grid areas (Fig. 2), record. Note if you see a lot of blue cells in any of your samples (~ number?) or if you see anything else that seems different.

17) Once you have compiled the data for all of your wells and all time points you will share this data with your research team so that each team member will have the data for both FBS concentrations and both of the control media at all 3 time points.

18) Each student should the using this data to generate a table with EXCEL (or google spreadsheets, if you prefer) and calculate the mean cell count (MEAN in EXCEL), and standard deviation (STDEV in EXCEL) for the control and each experimental cell population at each timepoint. Then generate a line graph of this date that includes error bars representing the standard deviation in your counts for each condition at each timepoint.

# ***Turn in the file containing your table and graph along with a short write up answering the following questions before the next lab session: What was the hypothesis your research team was testing? Does your data support your hypothesis? Why or why not?***

*Note: Assignments (graphs,write ups) should be turned in via the Lab Assignment on the course Moodle site.*