Advanced Light Microscopy Facility

Latest news

2023 Rates in effect for the SMOC (01-03-2023 to 29-02-2024)
Informative Flyer on Equipment available at SMOC
Informative Flyer on Super Resolution Informational at SMOC

Scientific supervisor: Francisco Javier Díez-Guerra

Technical director: Álvaro Sahún

smoa@cbm.csic.es

911 964 643

Third floor - 310

Presentation

Since 1999, the Advanced Optical Microscopy Facility (SMOA) has been in charge of the maintenance and management of the use of the advanced optical microscopy systems of the Centro de Biología Molecular Severo Ochoa (CBM), with a base of more than 300 annual users belonging to the CBM, as well as to other public and private organizations.

Our facility offers advice, support and training through the organization of theoretical-practical seminars on advanced microscopy techniques and image analysis.

Likewise, it is also in charge of the distribution of reagents and consumables related to microscopy, and the search for resources to acquire new systems that meets the demands of the different users of the CBM.

As a central scientific facility in the CBM, although it does not carry out any scientific research work, the SMOC has the following systems:

6 Laser scanning confocal microscopy (LSM) systems.
1 Spinning disk confocal microscopy system (SDCM).
1 Confocal microscopy system with a super-resolution stimulated emission depletion module (STED) and a fluorescence lifetime imaging module (FLIM).
3 Widefield microscopy systems with fluorescence that allow live cell imaging experiments and also fixed samples. And another Widefield microscope with a color camera for immunohistochemical samples.
In addition, it has 2 workstations for image analysis, a vibratome and a stereomicroscope.
An on-line system for accessing and managing system bookings and invoicing.
A stock of reagents and consumables related to microscopy.

In adition, our service is integrated into the Spanish Network of Advanced Optical Microscopy (REMOA) and has been a member of the former Network of Laboratories and Infrastructures (RedLab) of the Region of Madrid (CM) (Laboratory nº 216) since 2007.

Finally, since March 2009, our service has been quality certified according to the ISO9001:2015 standard issued by the company AENOR, which demonstrates SMOA's commitment to rigour and quality.

* Para llamadas desde el exterior a la extension xxxx se debe marcar: 34 91196xxxx
Apellidos	Nombre	Laboratorio	Ext.*	Horario	e-mail	Categoría profesional
Calvo Cazalilla	Elena	310	4643	9:30 - 17:00	elena.calvo@cbm.csic.es	Técnico Sup. de Actividades Técnicas y Profesionales. GP3
Gallego García	Carlos	310	4643	8:00 - 15:30	cgallego(at)cbm.csic.es	Titulado Sup. de Actividades Técnicas y Profesionales. M3
Sahún Español	Álvaro	310	4643	8:30 - 16:00	alvaro.sahun(at)cbm.csic.es	Titulado Superior FC1
Sánchez Jiménez	Carmen	310	4643	9:30-17:00	csjimenez(at)cbm.csic.es	Titulado Sup. de Actividades Técnicas y Profesionales. M3
Villalba Villacorta	María Teresa	310	4643	7:30 - 15:00	tvillalba(at)cbm.csic.es	Ayudante Investigación

STED & FLIM Microscope

STELLARIS 8 STED + FLIM

Location: 3^rd Floor (Lab. 310)

Confocal STELLARIS 8 with STED and FALCON (FLIM) coupled to an inverted DMi8 microscope (Leica)

Booking

Technical Specifications

Confocal Microscopes

Widefield Microscopes

Workstations for image analysis

Additional Equipment

Special Objectives

Deconvolution and calibration parameters for the SMOA equipment

Available reagents at the Advanced Light Microscopy Facility

Available reagents at the Advanced Light Microscopy Facility

Available reagents at the Flow Citometry Facility

Spectral viewers for fluorophore and fluorescent protein selection

Rates

2023 Rates (01-03-2023 to 29-02-2024)

General recommendations - Tips

Informative Flyer on Equipment available at SMOA

Informative Flyer on Equipment available at SMOA

Fluorophores and fluorescent proteins

Confocal or widefield?

Objectives, immersion oil and sample preparation

Controls for immunofluorescences

Colocalization experiments

Fluorescence quantification

Live cell imaging

Protocols

Autofluorescence

Cellular components that generate autofluorescence

Methods for removing autofluorescence

Sodium Borohydride (NaBH₄)
- Treat the fixed coverslips with NaBH4 (1 mg/ml in PBS, pH 8.0) for 10 minutes at room temperature (RT)
- Wash with PBS and continue with the immunofluorescence protocol. This solution must be recently prepared, observing bubble formation
Toluidine blue (‎C₁₅H₁₆N₃ClS)
In cases that FITC is the detection method and mainly for tissues. This will decrease autofluorescence mainly due to chlorophyll and cell walls (elastin/collagen):
- Incubate the tissues with 0.1% Toluidine Blue (Sakai, 1973. Stain Technology 48: 247-249). Incubation times could vary from several hours to overnight
- Wash off the solution excess
Ammonium chloride (NH₄Cl)
- Treat the fixed coverslips with NH4Cl (50mM diluted in PBS, pH 8.0) for 10 minutes at RT
- Wash with PBS and continue with the immunofluorescence protocol
Sudan Black (C₂₉H₂₄N₆)
- Treat the coverslips after secondary antibodies incubation with 0.3% Sudan Black (w/v) in 70% EtOH (v/v) for 10 minutes
- Wash extensively with PBS and proceed mounting the samples
Trypan Blue (C₃₄H₂₈N₆O₁₄S₄)
- Treat the coverslips after secondary antibodies incubation with 250µg/ml Trypan Blue in PBS pH 4.4, for 1 minute
- Wash extensively with PBS and proceed mounting the samples

Commercial products

Papers

ARTÍCULO	FUENTE
"Characterizing and Diminishing Autofluorescence in Formalin-fixed Paraffin-embedded Human Respiratory Tissue"	Journal of Histochemistry & Cytochemistry 2014, Vol. 62(6) 405–423
"Simple Method for Reduction of Autofluorescence in Fluorescence Microscopy"	The Journal of Histochemistry & Cytochemistry 2002, Vol.: 50 issue: 3, page(s): 437-439
"Reduction of Lipofuscin-like Autofluorescence in Fluorescently Labeled Tissue"	The Journal of Histochemistry & Cytochemistry 1999, Vol. 47(6): 719–730

Fixing

Paraformaldehyde 4% (PFA 4%)

Paraformaldehyde is a good fixation method for immunofluorescence assays. Although this method does not conserve the structural integrity of the cell as well as glutaraldehyde, the antegenic sites remains intact, this fixing method also does not generate too much autofluorescence.

Preparation:
1. Dissolve 4% paraformaldehyde (= 10% formalin) in distilled water preheated to 55-60ºC
2. Add some drops of 1M NaOH and shake until completely dissolved
3. Cool to room temperature (RT). Add PBS up to 1X and bring the pH to 7.4
4. Do not keep this solution more tan a week at RT (it can be conserved frozen in aliquots at -20ºC for several months)

Use:
1. Wash the coverslips with PBS and fix them with PFA 4% added slowly, for 20-30’
2. Wash the coverslips with PBS (3x5’)
3. Keep the fixed coverslips at 4ºC in PBS with 0.01% Azide or start immunofluorescence assay

Formaldehyde-Acetone

Use:
1. Wash the coverslips with PBS at 37ºC
2. Fix the cells with 4% PFA in PBS for 10’ at RT
3. Wash the cells with PBS (3x5’)
4. Permeabilize the cells with acetone at -20ºC for 10’. Incubate at -20ºC
5. Next: a) Dry the coverslips and keep them at 4ºC or b) rehydrate them with PBS for 10’ at RT, and start the immunofluorescence assay

Formalin (Formalin solution, Sigma- HT501128)

Use:
1. Wash the coverslips with PBS and fix for 20 'with formalin adding it slowly
2. Wash the coverslips with PBS (3x5’)
3. Keep the fixed coverslips at 4ºC in PBS with 0.01% Azide or start immunofluorescence assay

Methanol

Methanol is a good fixation method if we want to conserve the structural integrity of the cell, although the antigenic sites of binding to the antibody can be damaged. In addition, for effective permeabilization it may be necessary to use an anionic detergent.

Use:
1. Wash the coverslips with PBS
2. Add methanol at -20ºC slowly over the cells and incubate at -20ºC for 15’
3. Remove the methanol, rehydrate by washing with PBS (2x5’) and start immunofluorescence assay directly with the primary antibody incubation

Acetone

With this fixation method, the antigenic antibody-binding sites are very well preserved and the amount of background fluorescence generated is very low. However, the structural integrity of the cell disappears, although when visualized in widefield the structure remains sufficiently intact to analyze the subcellular location of the protein of interest. However, when an attempt is made to make a 3D study using confocal microscopy, it is appreciated that the cells fixed by acetone have lost their three-dimensionality. The cell structure, unlike fixation by methanol, is not visible through transmitted light. Acetone is a good permeating agent so subsequent detergent treatment is not necessary.

Use:
1. Wash the coverslips with PBS
2. Add methanol at -20ºC slowly over the cells and incubate at -20ºC for 10’
3. Wash the cells with PBS (3x5’)

Acetone-Methanol

Using an acetone-methanol combination is due to a compromise between the destructive nature of acetone and the lack of labelling from methanol. Alhough it is recommended to look for the best ratio, sometimes, it is a good idea to start with a 50:50 ratio. In some cases, the best ratio is 20:80 (acetone:methanol).

Use:
1. Wash the coverslips with PBS
2. Add acetone:methanol at -20ºC slowly over the cells and incubate at -20ºC for 15’
3. Remove the mixture and rehydrate by washing with PBS (2x5’). Start the immunofluorescence assay with the primary antibody incubation

Glutaraldehyde

Glutaraldehyde is the standard fixation method for electron microscopy. It preserves the structure very well, but unfortunately destroys most of the antibody-binding sites, generating, in addition, much autofluorescence in fixed cells. Incubating fixed cells with sodium borohydride solution for several minutes can decrease the amount of autofluorescence that is generated during the fixation with glutaraldehyde. A good solution, may be using a combination of 0.25% glutaraldehyde with paraformaldehyde.

Inmunofluorescence

Essential and Mandatory Controls in Microscopy:

Autofluorescence: Performing the same protocol with no primary or secondary antibodies.
Secondary antibodies: Staining the sample only with each one of the secondary antibodies.
Crosstalk: Staining with each of the primary and secondary antibody combinations separately to image all channels with the same acquisition parameters as used in double or triple preparations.
Simultaneous incubation versus sequential incubation: Although the simultaneous incubation in the same step with all the primary antibodies and then all the secondary antibodies is faster, it is more convenient to first incubate sequentially with one of the primaries, then its corresponding secondary, and so on.

Cell cultures

If necessary, remove autofluorescence.
Treat the coverslips with 0.1% Triton X-100 in PBS for 10 minutes at room temperature (RT).
Incubate the coverslips with Blocking solution (1-5% BSA or 1-5% Foetal Calf Serum (FCS) + 0.1% Triton X-100 in PBS), or MAXblock™ Blocking Medium (Active Motif), for 10 minutes at RT.
Incubate the cells with the primary antibody diluted in Blocking solution for 1h at RT. In order to remove aggregates and/or reducing background staining, centrifuge the antibodies in a minifugue at 12.000xg for 5 minutes at 4°C, previously.
Wash the coverslips by submerging them in PBS 3 times.
Incubate the cells with the secondary antibody diluted in Blocking solution for 45 minutes at RT and darkness. In order to remove aggregates and/or reducing background staining, centrifuge the antibodies in a minifugue at 12.000xg for 10 minutes at 4°C, previously.

NOTE. In case of using phalloidins, it is better to prepare the secondary dilution in PBS only.
Wash the coverslips by submerging them in PBS 3 times.
Submerge the coverslips in distilled water once, and blot excess on paper.
Pass the coverslips briefly in EtOH and blot excess on paper (This step is not strictly necessary).
Immediately afterwards mount the coverslips on slides.
Sample MOUNTING:
- With a yellow tip, place a drop of mounting medium on the slides.
- Then gently place the coverslip with the cells face-down on the drop of mounting medium and press lightly with filter paper so that only a thin layer of mounting medium remains between the slide and the cover.
  
  IMPORTANT. In case of very thick preparations, be careful with the previous step because the real volume of the simple could be altered.
- Let dry for 24h at RT and darkness on filter paper. Samples can be dried in an oven at 37°C for 3-4h, but preferably at RT.
- Seal the edges with nail polish. Only if the preparations will be stored for a long time.
- Stored at 4°C in the dark they can last for many months. Never store the preparation at 4°C before it is completely dry.

Tissues

General tissue protocol

NOTE: If possible, carry out the following steps with floating sections and on a rotary shaker.

Use some method to remove autofluorescence.
Incubate with the permeabilization solution (0.1% Triton X-100 in PBS) for 15 minutes at room temperature (RT).
Incubate with blocking solution (1% FCS + 0.1% Triton X-100 in PBS) for 1-2 hours at RT.
Incubate with the primary antibody diluted in the blocking solution, overnight at 4°C. Centrifuge it previously at 12,000xg in the minifuge for 10 minutes at 4°C. For sections larger than 50 microns it is possible to extend the incubation for two nights by doing it at RT. In this case, add 0.02% Sodium Azide.
Wash 5 times for 5 minutes with PBS.
Incubate with the secondary antibody diluted in the blocking solution for 12-18 hours at RT in the dark. Centrifuge previously at 12,000xg in the minifuge for 10 minutes at 4°C.
Wash with the permeabilization solution for 1 hour with frequent changes.
LWash with PBS for another hour with frequent changes. The sections can be stored in PBS or 4% Paraformaldehyde (PFA) for several days. It is preferable not to go to the next step until they are going to be observed.
Put the sections on slides, add mounting medium and place a cover on top.

Tissue protocol with DAB

Nervous tissue protocol

Protocol elaborated from “Unraveling human adult hippocampal neurogenesis”. Nat Protoc 15, 668–693 (2020).

NOTE: The following conditions are suitable for particular primary antibodies and 50 µm human brain sections. Try different incubation times to set the antibodies of interest.

Fixation: Immediately after extracting and dissecting the tissue, immerse it in 4% PFA for 24h at 4 ° C. Wash 3 times for 30 seconds with 0.1N PB (phosphate buffer) at room temperature (RT).
Embed the tissue in agarose:
- Prepare a solution of 10% sucrose-4% low-melting point agarose, heated in the microwave to make it liquid and fill the well of a M6-M12 plate.
- Put the plate on ice and immerse the tissue. It may float for the first 2 minutes. Push it to the bottom to avoid it and wait for the agarose to completely solidify.
- Cut a cube containing the tissue, with 2-3 mm extra agarose in all dimensions.
Sectioning: Cut 50 µm sections on the vibratome, collecting the sections in 0.1 N PB.
Incubate the sections with 0.5% NaBH4 in 0.1N PB for 30 minutes (this step is not necessary if the tissue comes from mice). Put the plate at RT on an orbital shaker with gentle shaking. The appearance of bubbles is frequent.
Wash 5 times for 30 seconds with PB-Triton-BSA (PBT-BSA).
Antigen retrieval (antigen retrieval) with citrate (HC-AR), if necessary:
- Fill 10 mL glass vials with 5 mL of 1x Citrate buffer (pH 6.0) preheated at 90 ° C (make the buffer just before using).
- Place 1 to 4 sections belonging to the same type of sample in each glass vial, and close without sealing it to avoid a pressure increase during microwave heating.
- Subject the vials to 5-6 cycles of 10-20 seconds in a microwave (800W at maximum power). To avoid tissue damage, the liquid should never be boiled, in which case microwave cycles should be suspended. Wait a few seconds between each microwave cycle.
  
  NOTE. This process should be repeated until the edges of the sections begin to curve slightly, stopping as soon as this effect occurs. Excessive folding of the edges of the sections should be avoided to avoid damaging the sample, the appearance of backgrounds and to preserve the specificity of the signal.
- Close the vials well and immerse them in a water bath at 80 ° C for 20 minutes.
- Leave the vials closed for 20 minutes at RT.
- Remove the sections from the vials with the help of a soft brush and wash them 5 times for 30 seconds in 0.1 N PB.
Incubate with the primary antibody. Put the primary antibody diluted in PBT-BSA in each well of an M24 plate, place the sections in the wells and make sure that they are floating (350-400 µl per well in this type of plate is suitable for incubating up to two sections). Incubate for 5 days at 4 ° C with gentle shaking. Incubation conditions and time must be adjusted experimentally for the antibodies used.
Wash sections 5 times for 30 seconds with PBT-BSA.
Incubation with the secondary antibody. Put the secondary antibody diluted in PBT-BSA in each well of an M24 plate (500 µl per well is suitable for up to three sections). Incubate in the dark overnight (ON) at 4 ° C with gentle shaking.

IMPORTANT. From this step the sections should be kept in the dark. For this, aluminium foil can be used.
Wash the sections three times for 30 seconds with 0.1 N PB.
Incubation with DAPI (optional). To stain the nuclei, the floating sections can be stained with DAPI (1: 5000 v/v in 0.1 N PB) for 10 min at RT with gentle shaking. A volume of 500 µl per well (M24) can be used to stain a maximum of three sections.
Wash sections 3 times for 30 seconds with 0.1 N PB at RT.
Autofluorescence removal:
- Immerse the sections in 1 mL of 70% ethanol for 5 minutes with gentle shaking. In an M24 a volume of 500 µl is sufficient for a maximum of three sections per well.
- Incubate sections in commercial Autofluorescence Eliminator Reagent (EMD Millipore, Cat#2160) (Sudan Black diluted in ethanol) for 5 minutes with vigorous shaking to prevent reagent precipitation. In an M24 a volume of 200-300 µl should be sufficient (take in mind that the sections will be stained black). This treatment should be adjusted according to the native autofluorescence of the tissue.
- Wash the sections 3 times for 1 minute with 1 mL of 70% EtOH with strong shaking. Make sure that the Autofluorescence Eliminator precipitates are removed with these washes.
Wash the sections 3 times for 30 seconds in 1 mL of 0.1 N PB at RT with gentle agitation.
Mounting the sections. Place the sections on a slide treated with 2% gelatine. The use of a commercial mounting medium or not is optional. Once the mounting medium has been added, place a coverslip and allow the preparation to dry, protecting it from light. Wipe off excess mounting medium carefully.
Samples can be stored at RT in the dark. It is recommended to acquire the images within the month following assembly.

USED SOLUTIONS

10% sacarose-4% low-melting agarose (500 mL)
- Heat 500 mL of 0.1 N PB.
- Add 50 g of sucrose and mix on a magnetic stirrer.
- Once the sucrose has completely dissolved, add 20 g of Low Melting Point agarose. Warm in a microwave until bubbles begin to form. Shake and rewarm. Repeat until transparent and slimy.
  
  NOTE. The solution can be stored at -20 ° C for 6 months or at 4 ° C for a week.
0.1 N PB Solution (1 L)
- Add 14 g of K₂HPO₄ to 800 mL of bidistilled water.
- Once completely dissolved, add 2.65 g of NaH₂PO₄-H₂O.
- Adjust the pH to 7.4 and make up to 1L with distilled water.
  
  NOTE. It can be prepared and stored at RT for up to 6 months.
PB-Triton-BSA Solution (PBT-BSA) (100 mL)
- Dissolve 1 g of BSA (1% w/v) in 100mL of 0.1 N PB.
- Add 1 mL of Triton-X-100 (1% v/v) and mix on a magnetic mixer until completely dissolved.
  
  NOTE. The PBT-BSA solution can be stored at 4°C for one week
2% gelatine-coated glass slides
- Dissolve 6 g of porcine gelatine in 300mL of hot bidistilled water on a magnetic mixer.
- When completely dissolved, add 0.5 g of KCr(SO₄)₂.
- Pour 250 mL of this solution into the slide cell.
- Place 10 slides in the holder and immerse them in the gelatine solution for 1 min.
- Allow slides to dry ON before use.
  
  NOTE. These slides can be stored at RT for 6 months.

Adult drosophila gonads (ovaries and testicles) protocol

Links

Mounting media

Test different mounting media because it may happen that the signal of some channel diminishes or even disappears simply by using a media incompatible with the staining we are carrying out.
For 3D analysis, the mounting medium used must not harden.

Mowiol/Dabco

Mix 6 g. of glycerol with 2.4 g. of Mowiol (Calbiochem Ref.475904, Polysciences Ref.17951) in a 50 ml centrifuge tube with a magnetic stir bar.
While mixing, add 6 ml of MiliQ water and leave for 2 hours at room temperature (RT).
Add 12 ml of Tris 0.2M, pH 8.5 and 230 μl of Thimerosal 1% (w / v in water) (antibacterial and antifungal agent).
Incubate in a 50 ° C bath for 10' with frequent agitation to dissolve the Mowiol. This operation should be repeated until the Mowiol is completely dissolved.
Add 2.5% DABCO (1,4-diazobicyclo- (2,2,2) -octane).
Centrifuge at 5000xg for 15' to remove fines. Store in 1 ml aliquots at –20ºC. It is stable for one year. After using, store at 4ºC (until one month). Discard it when crystalline deposits begin to appear in the preparations.

Propyl Gallate

Phenylenediamine

PVA-Dabco

Murray's Medium

Commercial products

Coverslip treatment and adhesion

Live-cell imaging

Microscopy techniques

Software and Books

Software

Books

Manuals and Seminars SMOA

Videos of image analysis/treatment SMOA

Macro task automation

Macro Example

División de Canales

Esta macro permite guardar todos los canales de una imagen por separado y una imagen de combinación de todos ellos.

MACRO IMAGEN DE PRUEBA

FUNCIONAMIENTO DE LA MACRO:

prueba.tif

1º Seleccionar la carpeta donde están las imágenes para contar.

2º indicar la carpeta donde se guardarán los resultados (Se crea una nueva carpeta “Resultados”).

3º Indicar el formato de las imágenes (tif, lif, lsm, czi, nd2, vsi, ics).

4º Seleccionar el color para cada canal.

Nota: Si alguno de los parámetros no se introducen correctamente la macro lanzará un error y se parará.

Pasos del tratamiento:

Abre la imagen y separa los canales.
Guarda la combinación y los canales por separado en la carpeta “Resultados” con el prefijo (“Merge” o “ChannelNº”) y el nombre original.

Resultados obtenidos:

Una imagen de cada uno de los canales por separado en formato RGB.
Una imagen de Composite (mezcla de todos los canales).

1234

Contaje

Cuantificación de la Intensidad

Deconvolution

Advanced Light Microscopy Facility (Lab 310)

Centro de Biología Molecular Severo Ochoa (CBM), (CSIC-UAM)

C/Nicolás Cabrera,1

Universidad Autónoma de Madrid. Cantoblanco.

28049. Madrid

Teléfonos:

Despacho: 91 196 4613

Laboratorio 310: 91 196 4643

Laboratorio 336: 91 196 4660

Fax. 91 196 4420

E-mail: smoa@cbm.csic.es

SMOC en 3 plantab 1

Transporte en la Comunidad de Madrid

Transporte en la U. Autónoma de Madrid

RENFE

Líneas C7, C8 y C10 de Cercanías: Parla-Atocha-Chamartín-Cantoblanco-Alcobendas/S.S. de los Reyes o Colmenar Viejo.

Estación de Cantoblanco.

AUTOBUSES INTERURBANOS

Líneas 714, 827, 827B y 828

METRO DE MADRID Y LÍNEAS DE AUTOBUSES EMT

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Curing (hard-set)	ProLong Glass, Diamond, Gold. (ThermoFisher Scientific)
Curing (hard-set)	Vectashied Vibrance (Vector Laboratories).
Aqueous (non-hardening)	Vectashield (Vector Laboratories)
	Fluoromount G (ThermoFisher Scientific)
	Glycerol (several companies). 50% (DIC), 75% (DIC+IFI), 90% (IFI)

CENTRO DE BIOLOGÍA MOLECULAR SEVERO OCHOA

Advanced Light Microscopy Facility

STELLARIS 8 STED + FLIM

LSM900 Upright Confocal Microscope

Spinning Disk SpinSR10 Confocal Microscope

LSM800 Inverted Confocal Microscope

In vivo Nikon A1R+ Confocal Microscope

LSM710 Upright Confocal Microscope

LSM710 Inverted Confocal Microscope

Orca Flash (sCMOS-monochrome)

CMOS-color Camera

PCO (FRET)

High-speed in vivo system

Orca-Fusion

(still in development)

Workstation 1 (Metamorph)

Workstation 2 (Deconvolution)

Stereo Microscope with Cold Light (Leica)

VT1200S Vibratome (Leica)

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