Available laser lines and corresponding fluorophores

Dyes labeled * should be avoided – the signal is low and they photobleach easily. Instead, use modern fluorophores if possible, such as DyLight, Chromeo, Alexa Fluor, Abberior, Atto, PromoFluor, or Cyanine dyes (Cy3,Cy5, ..) and mount the sample in a compatible anti-fade mounting medium.
** Useable, but excitation is not ideal (laser line too far away from the absorption peak)

Laser, Emission wavelength	Fluorophores
405 nm	DAPI, Hoechst, Nuclear Blue
Argon, 458 nm	CFP, mTFP, mCerulean and similar cyan-emitting proteins
Argon, 488 nm	GFP, mNeonGreen, Alexa Fluor 488, DyLight 488, FITC*, Fluorescein*
Argon, 515 nm	YFP, mVenus, mCitrine
HeNe, 534 nm	mCherry, tdTomato, dsRed, Cy3, Alexa Fluor 568, Chromeo 546, Alexa Fluor 594**
HeNe 633 nm	Cy5, Alexa Fluor 633, Alexa Fluor 647, Atto 647N, …

Fluorophore combination for multi-channel imaging

Choose dyes with well separated excitation and emission peaks. A combination of dyes with overlapping spectra is NOT recommended (e.g., FITC and Cy3). When staining for DNA with DAPI or Hoechst dyes, remember that they have broad emission spectrum and will have an overlap with green-emitting dyes like FITC, Alexa Fluor 488. Sequential imaging will be necessary.

*There is no dichroic mirror set for simultaneous 405nm and 633 nm excitation. The Dye Assistant in the software will in such case select a 20/80 beamsplitter – this is workable but not ideal. There is a work-around – please ask MIC staff to show you how to set the imaging conditions.

Number for Fluorophore	Suggested dyes
DNA + 1 additional channel	DNA: DAPI or Hoechst (405 nm laser) 2nd channel: Cy3/Alexa 568 (543nm laser) or Cy5/Alexa 647* (633 nm laser) Alternative 2nd channel: EGFP/Alexa Fluor488 (488 nm laser) – use sequential imaging
DNA + 2 Additional channels	DNA: DAPI or Hoechst 2nd channel: Cy3/Alexa 568 (543nm laser) 3rd channel Cy5/Alexa 647* (633 nm laser)
DNA + 3 additional channels (requires Virtual Channel Scan setting)	DNA: DAPI or Hoechst 2nd channel: EGFP/Alexa 488 3rd channel : Texas red or Alexa 594 4rd channel Cy5/Alexa 647
two fluorescent proteins	EGFP/mNeonGreen and mCherry/tdTomato (use the GFP/RFP filter cube and external HSDs for weak signals) CFP and YFP (use the CFP/YFP filter cube and the external HSDs for weak signals)

Mounting Media

Best image quality is achieved if the mounting medium has the same refractive index as the immersion medium for the objective (Water – 1.33, Glycerol – 1.47, Oil – 1.518). To reduce photobleaching of the fluorophores, the the mounting media may contain anti-fade reagents. Some fluorophores are not compatible with certain antifade reagents. Consult the mounting media datasheet for compatibility. Here are some standard mounting media and their refractive indices:.

The refractive index of your mounting media should match as closely as possible the refractive index of your imaging objective’s immersion medium

• 75% Glycerol – 1.44
• Citifluor 1.463
• Vectashield – 1.457
• Mowiol – 1.49
• ProLong Gold (hardening version) – Fresh mounting: 1.39 Cured for 1 day: 1.40 Cured for 4 days: 1.44

None of these media are an exact match for the oil immersion or water immersion objectives. However, the Leica SP8 confocal system in the MIC is equipped with a multi-immersion 20x objective (adjustable for water, glycerol, oil immersion) and also a high resolution 63x/1.3 glycerol immersion objective). For the Olympus microscope, the mounting media should be chosen based on which microscope objective is going to be used for imaging (water immersion 60x/1.2, or the oil immersion objectives).

For more information, see this paper: Ravikumar S, Surekha R, Thavarajah R. Mounting media: An overview. J NTR Univ Health Sci 2014;3, Suppl S1:1-8

• Live samples, or samples mounted in a buffer are best imaged with a water immersion objective. Using oil immersion objectives for these samples is NOT recommended, because resolution and signal strength will degrade rapidly when focusing deeper into the sample.
• For routine imaging of fixed, fluorescently stained samples, non-hardening glycerol-based mounting media are convenient and work reasonably well with oil immersion objectives. You can make your own media (see links to recipes below) or use commercially available products like Vectashield, Prolong Gold, .Citifluor AF1, …
• For highest resolution imaging with oil immersion objectives, the samples should be mounted in a medium that has the same refractive index as glass and immersion oil (1.515). This will minimize spherical aberration. Commercial media include Citifluor CFM, and Prolong Glass. If it is a hardening medium, it may shrink during curing and distort 3D samples. The 2,2- thiodiethanol is a suitable high-index medium, but is not compatible with fluorescent proteins at the optimal (97%) concentration. T. Staudt, M. C. Lang, R. Medda, J. Engelhardt, S. W. Hell, 2,2′-thiodiethanol: a new water soluble mounting medium for high resolution optical microscopy. Microsc. Res. Techn. 70, 1-9 (2007).

Recipes for Anti-fade Mounting Media

1. Medium with n-propyl gallate. From Jackson Immunoresearch website www.jacksonimmuno.com: “In our experience n-propyl gallate added to mounting media reduces fading of fluorescence from many different fluorophores during fluorescence microscopy. The following is a simple recipe for making an anti-fade mounting medium containing n-propyl gallate.”

• Prepare a 10X PBS stock solution.
• Prepare a stock solution of 20%(w/v) n-propyl gallate (Sigma P3130) in dimethyl formamide or dimethyl sulfoxide. (Note: n-propyl gallate does not dissolve well in water-based solutions.)
• Thoroughly mix 1 part of 10X PBS with 9 parts of glycerol (ACS grade 99-100% purity) and slowly add 0.1 part 20% n-propyl gallate dropwise with rapid stirring. 

Store the medium in aliquots at -20 C.

2. Medium with p-phenylenediamine (from CSHL Protocols http://cshprotocols.cshlp.org/content/2019/2/pdb.rec104265.full?rss=1):

Reagent	Amount to add	Final concentration
Tris-HCl (1 M, pH 8.8)	0.8 mL	20 mM
p-phenylenediamine	0.20 g	0.5%
Glycerol (100%)	36 mL	90%

Mix the above ingredients together in a 50-mL conical tube. Adjust volume to 40 mL with H2O. Dissolve the solids by slowly bubbling nitrogen gas through a Pasteur pipette inserted deep into the mixture. Filter through a 0.8-µm filter at room temperature. Store at −20°C in 1- or 3-mL syringes devoid of air bubbles. Discard when dark brown. Handle this reagent with gloves; it causes stains when spilled.

Imaging Chambers, Slides and Plates

Microscope objectives are designed to view the sample through a thin glass window, 0.17 mm thick (this is the #1.5 coverglass). On the inverted microscope, the bottom of the sample chamber must be thin and optically clear. If your samples are in a standard plastic Petri dish or a regular multi-well culture plate, you will not be able to get the best image quality with the 10x objective, and you will not be able to focus at all with the other objectives.

There are two long-working dry objectives, 20x/0.4 and 60x/0.7, that may be used with these samples. These objectives are NOT installed on the microscope. Please consult MIC staff for specific instructions.

Recommended types of samples:

• Slides with a 0.17 mm thick coverglass, sealed with nail polish or other suitable sealer that has dried
• Chambered slides where the culture chamber was removed after staining, suitable mounting medium and a coverglass was applied.
• Chambered slides with optically clear, thin bottom (e.g. Ibidi microslides, Greiner CELLview cell culture Slides)
• Imaging-type 96-well plates with optically clear, thin bottom – these MUST be specifically the type where the bottom is minimally recessed relative to the skirt of the plate. For example, Greiner Screenstar microplate (Item No.: 655866) or Eppendorf Cell Imaging Plate (Cat. # 003074103), or Ibidi µ-Plates should work well. CONSULT THE MIC STAFF IF YOU ARE NOT SURE WHICH PLATE YOU NEED.

NOT Recommended:

• LabTek II chambered coverglass – the chambers are too close to the end of the coverglass, pose a risk of damaging microscope objectives
• 96 well plates with deeply recessed bottom – They create a risk of damaging microscope objectives. The edge rows and columns CANNOT be imaged,
• If you use these plates or chambers, when the objective is in focus and you move to the well on the end or edge of the plate, the metal sample holder will hit the objective and damage it. You and your PI will be responsible for objective repair/replacement ($$$).