DNA ploidy and genome size estimation using flow cytometry (Dan B.)

Flow cytometry (FCM) is – in principle – a relatively simple technique that allows one to measure properties of cells or organelles as they are intercepted (individually) by a high-intensity light source. In plant biology, it is often used to analyze extracts of intact nuclei, with the aim of determining DNA content and (indirectly) ploidy levels (this is the application I will focus on here). A good reference (including troubleshooting tips) can be found here.

There are three main steps involved in a FCM analysis: sample preparation, data acquisition, and data analysis. Among these, sample preparation is by far the most crucial part. Basically, if this step goes well, data acquisition and analysis are relatively straightforward.

I. Sample preparation

Note: all utensils needed for this step are in the lab, in the cupboard labeled “flow cytometry” (just under the big centrifuge). The reactives are on the chemical shelf.

Sample preparation involves mechanical homogenization of the tissue, and isolation of intact nuclei.

An important point to make here is that during FCM, you measure relative fluorescence intensities. Therefore, interpretation of results in terms of ploidy and genome size also requires the analysis of a standard (i.e. plant of known ploidy and/or genome size). We have in the lab seeds for a number of widely accepted standards for plant FCM:

Raphanus sativus (2C = 1.11 pg DNA)

Solanum lycopersicum (2C = 1.96 pg DNA)

Glycine max (2C = 2.50 pg DNA)

Zea mays (2C = 5.43 pg DNA)

Pisum sativum (2C = 9.09 pg DNA)

Secale cereale (2C = 16.19 pg DNA)

Vicia faba (2C = 26.90 pg DNA)

When choosing a reference standard for your species of interest, you want to make sure the genome of the standard does not differ from that of your species by more than twofold (as this can lead to errors due to nonlinearity if instrument calibration is poor). Also, you should avoid using a standard with very similar nuclear DNA content (due to potential overlap in FCM reads for sample and standard). For diploid Helianthus species I use Zea mays, while for tetraploids and hexaploids I use Vicia faba and Secale cereale, respectively.

There are various methods of standardization, including internal standardization (whereby sample and standard are co-chopped, and nuclei are isolated and analyzed concomitantly), or external standardization (whereby nuclei are isolated and analyzed independently for sample and standard). Generally, for problematic tissue with (potentially) high content of inhibitors (such as sunflower tissue), it is recommended that you use internal standardization (this way, the inhibitors should influence both standard and sample nuclei in a similar manner, minimizing potential error).

With regards to the type of tissue, you can use either fresh or silica-dried material. Although I cannot comment on the latter option (as I have not used it so far), genome size measurements are generally considered most reliable when performed using fresh tissue. Ideally, you want to use almost fully expanded leaves, sampled from intact, sufficiently watered plants. I generally collect the tissue and use it right away to isolate nuclei, although you can also store it in a Petri dish, wrapped in a wet paper towel, at 4°C for a few days. Eric Baack (another Riesebergler that used FCM for Helianthus and whom I contacted for advice) said freezing the tissue lead to complete failure, so I did not try that. I use 30-40 mg of tissue (for each of standard and sample), although you can increase this amount (if the yield of nuclei is not so good), or decrease it (if the content of inhibitors is too high).

Word of caution: avoid using very young leaves (as those may have a high content of inhibitors), as well as material colonized by pests (due to the obvious risk of biological contamination or accumulation of defense substances by the plant). Also, do not use senescent tissue as it may comprise endopolyploid nuclei.

The collected tissue should be chopped (for about 60 seconds) with a sharp razor blade (to release nuclei) in 1 mL of the isolation buffer, in a Petri dish. Use force, you want to chop straight through the material. Naturally, the amount of buffer will vary depending on whether you chose to increase or decrease the amount of tissue recommended above. If you do not use enough buffer, the tissue will stick to the blade (preventing proper chopping). If you use too much buffer, you will find yourself having to chase bits of tissue in the Petri dish. The chopping should (ideally) be performed on a cold surface (I use an aluminum block). The buffer should also be cold (I store it at 4°C).

An important note to make here is that the plant extract should remain green at all times. If you find that during chopping (or at any other point after that) it is turning yellow or brown, you know you have a problem with secondary metabolites (these phenolic substances often turn brown when oxidized). Such compounds may interfere with staining and/or exhibit autofluorescence. In that case, I suggest first looking into the quality of the tissue or buffer you are using.

Word of caution: Although seemingly trivial, the chopping part is just as important as any other step in sample prep. If you do not do it for long enough, you will not obtain too many nuclei. Conversely, if you do it for too long, you might overwhelm the buffering ability of the buffer.

The nuclear isolation buffer is another very important component of sample prep. It serves several important tasks, including to:

– facilitate the isolation of intact nuclei, free of adhering cytoplasmic debris

– maintain the stability of nuclei and prevent their aggregation

– protect the DNA from degradation

– provide an appropriate environment for specific staining of DNA

Although there are several “general purpose” buffers available, no single buffer works well with every plant species (due to the diversity of tissues in chemical composition and/or structure). Eric Baack successfully used a modified version of what is known as the Tris.MgCl2 buffer (reference is here). After trying this and several other buffer alternatives, I chose a slightly modified version of de Laat’s buffer, which gave me the best resolution FCM reads.

The recipe for this magic potion is:

15 mM HEPES (organic buffer that acts to stabilize the pH of the solution and to keep nuclei intact)

1mM EDTA (chelating agent used to bind divalent cations, which serve as nuclease cofactors)

0.2 % (v/v) Triton X-100 (non-ionic detergent, used to release and clean nuclei, and decrease the aggregation affinity of nuclei and debris).

80 mM KCl (inorganic salt used to achieve proper ionic strength)

20 mM NaCl (also an inorganic salt)

300 mM sucrose (used to provide isotony)

0.5 mM spermine (chromatin stabilizer)

15 mM β-mercaptoethanol (antioxidant that should keep phenolics in a reduced state)

0.25 mM PVP-40 (its amide group is available for binding with the cytoplasmic inhibitors, and so it competes with proteins and DNA in interacting with these compounds).

Adjust the pH of this buffer to 7. It is important to know that the staining with the fluorochrome (detailed below) is carried out at neutral or slightly basic pH. At lower pH values, other components (such as cytoplasm, RNA) are easily stained along with chromatin.

After chopping, the crude homogenate should be sieved through two layers of Miracloth (also found in the cupboard labeled “flow cytometry” in the lab) to remove any large particles which may clog the flow cytometer. The sieved extract should be collected in an eppendorf tube for the next step.

At this stage, you should have a clear suspension (without precipitate or tissue fragments) which can be of various shades of green (depending on the plant species used and the amount of tissue), containing nuclei but also other cytoplasmic constituents and debris. To isolate only (or mostly) nuclei, you use a centrifugation step. I generally use 5 min at 1200 RPM (note the reduced speed). After this step, you should see a white (!not green) pellet. Discard the supernatant and you are ready for the next step. If you do not see a pellet at all, try increasing centrifugation speed. Alternatively, if the pellet is green, discard the supernatant, resuspend in 1 mL of nuclear isolation buffer, and redo the centrifugation step. Usually, the second time you will get a white pellet.

The next step is optional, and should be performed if you do not intend to finish the protocol all the way immediately. It involves resuspending the pellet and fixing the nuclei in 1 mL of ethanol: acetic acid (v/v 3:1) mix for 30 minutes or more. The fixed nuclei can be stored just fine for several weeks at 4°C (reference is here). Apart from the convenience of being able to store the nuclei, this method (by involving an additional resuspention/centrifugation step) might help get rid of additional cytoplasmic debris. When you are ready to continue with the protocol, centrifuge the fixed nuclei under the same conditions as above, and discard the ethanol: acetic acid mix.

Whether you chose to use the optional step detailed above or not, you should have a white pellet for this next stage. Resuspend this pellet in 300 μL of nuclear isolation buffer. If, during data acquisition, you find the sample is too dilute (i.e. you are only measuring less than 50 nuclei per second), you can reduce this volume to 200 μL.

The next step is RNA-se treatment. This step is optional, depending on what type of dye you use. Ethidium bromide and propidium iodide (PI; this is what I use) intercalate into double-stranded DNA and double-stranded RNA. These dyes are suitable for estimation of DNA content in absolute units provided that, prior to staining, RNA is removed with RNA-se. Other dyes (such as DAPI) bind AT bases (and therefore do not require RNA-se treatment), but are not recommended for estimation of DNA content (especially if differences in base composition are expected).

The last step of sample prep is the staining of the nuclei with the DNA-selective fluorochrome. This fluorochrome should bind stoichiometrically to DNA. For PI, concentrations of 50-150 mg /l are appropriate (I use 100). Generally, it is recommended that you do the staining for a few minutes to up to 1 hour (so once you are at this step you want to continue with data acquisition as soon as possible).

II. Data acquisition

During this step, you analyze the optical properties (fluorescence and light scatter) of the nuclei suspension you prepared in the previous step, using a flow cytometer.

There are several flow cytometers available on campus, some of which are housed on the first floor of the Biomedical Research Centre (BRC), on Health Sciences Mall. The person that manages them is Andy Johnson (andy@ubcflow.ca). Andy also periodically organizes (approximately 3-hour) workshops on the basic principles of flow cytometry (also including a hands-on course on how to operate the instruments), so these are useful to take. Flow cytometers may appear to be complex instruments, but all basically consist of three main components:

1) fluidics – this part brings the sample to the instrument, and separates the nuclei suspension in a stream of single particles, using hydrodynamic focusing.

2) optics – consists of a light source (most often a laser) and a focusing system

3) electronics – consists of photomultipliers, which take the photons that have been scattered or emitted (as the fluorescently labeled particles are intercepted by the laser) and put out electrons (which are converted to a digital value, recorded on the computer).

The BD FACSCalibur flow cytometer I have been using at the BRC is connected to a (quite ancient) Mac. To access it, you need to log in using the Rieseberg lab account (ID: Rieseberg lab; password: Loren). This account is also how our lab is being charged for using the machine (i.e. based how long you are logged in, by the hour – remember to log out once you are done).

There is a proprietary software you are supposed to use to interact with the machine, CellQuest. To connect CellQuest to the cytometer, you use Acquire > connect to cytometer. I have saved CellQuest projects for diploid and polyploid Helianthus data collection. You can access those by using Open > Users FACSCalibur > Rieseberg Lab > Dan Bock > Helianthus diploid (or tetraploid/hexaploid) data analysis. These projects will plot several parameters during data acquisition, including forward scatter (a measure of particle size) or side scatter (a measure of particle optical complexity), which should help you identify homogenous populations of particles (nuclei).

An important data acquisition option to use is “Counters” (Acquire > Counters). This will tell you the number of events measured by the machine per second. The BD FACSCalibur comes with three options for sample delivery rate (slow, medium and fast). I always run samples on slow, as excessive delivery rates (i.e. higher than 100 events per second) results in broadening of DNA peaks.

To collect the data, load your sample and use “Acquire”. Note: when the “Settings” option is on, you cannot save the data you acquire. Using “Settings”, adjusting the voltage of the photomultiplier so that the smallest DNA peak (corresponding to sample/standard G1 nuclear DNA content) is positioned on channel 200 (the highest resolution one). Once this is done, uncheck the “Settings” option, to be able to save the data you acquired. Make sure you measure at least 5,000 (10,000 is better) particles before saving the data and moving on to the next sample. This will usually take between 10-20 minutes, depending on the concentration of nuclei in your sample.

III. Data analysis

There are different software available to help you analyze flow cytometry data; I use FlowJo.

First, by performing what is called “gating”, you aim to exclude cell debris and clumped nuclei from the raw data. Basically, data points representing intact nuclei (at G1 and G2) for the sample and standard should be easily distinguishable from debris background, and should form a distinct population of particles with similar light scatter and fluorescence properties. For estimates of genome size and ploidy, you are interested in G1 nuclei, although by taking into account G2 nuclei as well (i.e. checking if those have double the DNA content), you can check if there are any problems with instrument linearity.

Note: after gating, you want to make sure you have at least 1000 nuclei for the relevant (i.e. G1) population of nuclei, although some publications report analyzing even more than that.

After gating, you want to check the quality of the gated data. Populations of nuclei are represented as Gaussian distributions of fluorescence intensities. The quality of data is estimated based on the width of such distributions, by calculating the peak CVs (CV% = SD of the peak / mean position of the peak x 100). Generally, CVs of DNA peaks ranging between 1-5% are considered acceptable quality. In FlowJo, you can easily insert a function (to calculate CV) for any gated population of particles.

Once you have confirmation on the quality of your data, you can go ahead and estimate the ploidy and/or genome size of your sample, which is straightforward:

For ploidy estimation: sample ploidy = standard ploidy x mean position of sample G1 peak / mean position of standard G1 peak

For genome size estimation: sample 2C value = standard 2C value x mean position of sample G1 peak / mean position of standard G1 peak