06-DAPI.Rmd

# | DAPI + Cell Counting {#DAPI}
<font size="1">**Created By:** Caitlin Casar on 2020-01-27 <br /> 
**Last updated:** 2020-02-13 </font>

## Sample prep

1. Prepare your cells by fixation for 30 minutes at room temperature. 
    <ul>
    <li>Add 0.1 ml 4% paraformaldehyde/PBS solution to a 2ml tube</li>
    <li>Add 0.9ml cell suspension</li>
    </ul>
    
<ul>   
```{r out.width = "250px", echo=FALSE, fig.cap='Fix your sample.'}
knitr::include_graphics("images/DAPI-1.png")
```
</ul>

2. Use sterile foreceps to place a glass microfiber Whatman filter on the filter mount. Wet the filter with filtered DI water and vacuum so that it is damp.

3. Use sterile forceps to place a polycarbonate Whatman filter on top of the glass microfiber filter. Wet the filter with filtered DI water and vacuum dry to flatten evenly. (Beware of the “taco” effect - the surface tension of the water on the polycarbonate filter may cause it to fold up on itself and it is very difficult to re-flatten). 

4. Place the filter column over the stack filters and clamp down. 

5. Add a few ml of filtered DI water to the column, then add your fixed cell suspension avoiding the sides of the column. The water helps to evenly disperse the cells across the filter. If you accidentally drop your cells on the side of the column, use filtered DI water to wash them down. 
<ul>
```{r out.width = "250px", echo=FALSE, fig.cap='Add your sample to the filter column.'}
knitr::include_graphics("images/DAPI-2.png")
```
</ul>
6. Vacuum the fluids through. Open the vacuum line on the flask before the next step or it may draw a vacuum and pull your dye through. 
<ul>
```{r out.width = "250px", echo=FALSE, fig.cap='Vacuum your sample through the filter.'}
knitr::include_graphics("images/DAPI-3.png")
```
</ul>
7. Dim the lights and close the curtain, DAPI is extremely photo-sensitive. Drip filtered DAPI down the side of the filter column so as not to disturb your cells on the filter. Add enough to completely cover the entire filter. 
<ul>
```{r out.width = "200px", echo=FALSE, fig.cap='Dye your sample with DAPI.'}
knitr::include_graphics("images/DAPI-4.png")
```
</ul>
8. Set a timer for ten minutes. After the timer goes off, vacuum the dye to dry the filter. 
<ul>
```{r out.width = "200px", echo=FALSE, fig.cap='Vacuum the dye through the filter.'}
knitr::include_graphics("images/DAPI-5.png")
```
</ul>
9. Add a drop of immersion oil to a glass slide. Use sterile forceps to place the filter onto the drop of oil. 

10. Add a drop of immersion oil on top of the filter. Do not touch the filter with the oil dropper or it may contaminate the oil. Place a glass coverslip over the filter, taking care to push out the bubbles. 
<ul>
```{r out.width = "250px", echo=FALSE, fig.cap='Prepare your slide.'}
knitr::include_graphics("images/DAPI-6.png")
```
</ul>
11. Add a drop of immersion oil to the top of the glass coverslip. You are now ready to place the slide on the microscope stage for use with the 100x oil objective. 


## Operating the Microscope


1. Remove the dust cover and turn on the power source, then power on the microscope. 

2. Power on the X-Cite lamp. Once the lamp is turned on, do not power off for at least 30 minutes. 

3. Open the Zen software. If you open the software before the microscope has been powered on, the software will try to communicate with the scope and get confused and will need to be rebooted. 

4. Turn on the DAPI RL (reflected light) 

5. Carefully lower the objective onto the drop of oil on your coverslip. When the objective makes contact with the oil, you will see a flash of light. 

6. Continue to lower the objective until you encounter the focal plane. The focal plane tends to be far down in the Z-direction. If you go too far in the Z-direction you will crack your slide and potentially damage the objective lense, so take your time! 

7. To view the image on the software, pull the eyepiece rod halfway out to direct 50% light to your eye and 50% to the camera. Pull the rod all the way out to direct light 100% to the camera. Click the “Live” button on the Locate tab in the laft panel and set the camera exposure. 

8. To acquire an image, click the “snap” button. If you want to save the images in their proprietary format, right click the image on the right panel and save. If you want them in jpg format, switch to the processing tab in the left panel and export the images to your file folder. 

9. You can add a scale bar with the graphics tool. 

10. When you’re finished, turn off the reflected light and remove your slide. Use lens paper to clean the oil off of the objective. 

11. Power off the software, then the microscope, then the power supply. Turn off the X-Cite lamp. 

12. Replace the dust cover on the microscope.


## Cell Counting

Bacteria densities (with the proper dilution) should be at least 30 organisms per field. 
<ul>
<li>Count at least 10 fields (to achieve a final count of 300 bacterial cells). </li>
<li>Calculate final bacterial densities using the following equation (from Wetzel and ikens, 1991).</li> 
</ul>
</font>


<font size="1">
Bacteria ml-1 = (membrane conversion factor * ND) </br>
Membrane conversion factor = Filtration area/area of micrometer field</br>
N = Total number of bacteria counted/number of micrometer fields counted </br>
D = Dilution factor; volume of sample stained/total volume of sample available 

<ul>
```{r out.width = "250px", echo=FALSE, fig.cap='Glowing cells on a filter.'}
knitr::include_graphics("images/DAPI-7.png")
```

```{r out.width = "250px", echo=FALSE, fig.cap='Count total of at least 300 cells, calculate average # cells/frame'}
knitr::include_graphics("images/DAPI-8.png")
```

```{r out.width = "250px", echo=FALSE, fig.cap='#cells/frame   x   #frames/filter  =  total cell count/volume'}
knitr::include_graphics("images/DAPI-9.png")
```
</ul>

## Troubleshooting

This is a step by step guide to troubleshooting issues that arise during microscopy. Many things can go wrong and it's important to approach it as a process of elimination. 

<font size="3">**1. Is something wrong with the microscope?**</font>

The microscope kit includes a DAPI test slide. Can you can see the specimen on the slide?
<ul>
<li>**Yes!** There is no technical issue with the microscope. Proceed to step 2.</li>
<li>**No!** Check that the eye piece shutter is open and the light path is directed 100% to the eye piece (not to the camera). Additionally, check that the Apotome filter is not blocking the light path. Once you've checked for these issues, if you still can't see the specimen it may be time to call a technician.</li>
</ul>

<font size="3">**2. Is something wrong with the antifade reagent?**</font>

The antifade reagent has a shelf life. When precipitates form in the solution, it alters the refraction index of the solution and can obscure your image. Prepare an overnight culture of E. coli - **do not use your own sample or an old culture of E. coli.** Why? Because your sample may have issues (for example, your cell density may be too low), and because unhealthy cells do not stain as well as healthy cells. 

Check that you can see cells by mounting 15 microliters of the culture on a slide and imaging via DIC. If you cannot see cells via DIC, your culture is not turbid enough. Wait until the culture becomes more turbid, then repeat this step before moving on to the next step.

Next, stain 100 microliters of the culture and mount the filter with Citifluor, then repeat but instead replace the Citifluor with immersion oil. **Make sure you do not have two coverslips stacked on top of your filter and be sure to add immersion oil on top of the coverslip - failing to do this will result in a blurry image!** Can you see cells on the filter without Citifluor but cannot see cells on the filter with Citifluor?
<ul>
<li>**Yes!** The Citifluor is the issue. </li>
<li>**No!** If you can't see cells on either filter, the Citifluor is not the issue. Proceed to step 3.</li>
</ul>

<font size="3">**3. Is something wrong with the dye?**</font>

The working solution of DAPI in the fridge has a shelf life of ~3 months due to exposure to oxygen and light. Prepare a fresh solution of DAPI from the freezer stock. The freezer stock concentration is 1mg/mL. Dilute 15 microliters of the stock solution in a 15ml falcon tube by topping to 15mL with filter-sterilized milliQ water for a final concentration of 1microgram/mL. 

Stain 100 microliters of E. coli with both the old and new dye. Can you see cells on the filter dyed with new dye but cannot see cells on the filter dyed with old dye?

<ul>
<li>**Yes!** The dye is the issue.</li>
<li>**No!** If you cannot see cells on either filter, the dye is not the issue. Proceed to step 4. 
</ul>

<font size="3">**4. What else could be wrong?**</font>

The most common issues are with the scope, Citifluor, and dye. If these are not the issues, there are a few other things to check - inspect the filter tower components for defects or filter residue on the frit that might be preventing a seal. Try increasing/decreasing the excitation lamp intensity - the higher your intensity, the quicker the dye will bleach and prevent you from seeing cells. Test a batch of dye that works in another lab - maybe our freezer stock has degraded (this happens within a couple years!).