Protocol to study adult neurogenesis in fresh-frozen human hippocampal tissue using an immunofluorescence quantitative approach

Summary

Adult hippocampal neurogenesis (AHN) results in the generation of new neurons throughout adulthood, thereby contributing to key hippocampal functions such as learning and mood regulation. Here, we present a protocol for the immunofluorescence-based detection of AHN markers in fresh-frozen human hippocampal sections. We describe steps for tissue fixation, antibody incubation, and autofluorescence elimination. We then detail procedures for image acquisition and quantification. This protocol allows the reliable identification of distinct cell populations and could be adapted for broader immunohistochemical applications.

For complete details on the use and execution of this protocol, please refer to Márquez-Valadez et al.¹

Graphical abstract

Highlights

• •Protocol for immunostaining fresh-frozen postmortem human hippocampal tissue
• •Reliable detection of markers of adult hippocampal neurogenesis in human tissue
• •Provides guidance for acquiring confocal images and determining cell densities
• •Guidance for morphometric determinations and dentate gyrus vasculature analysis

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Adult hippocampal neurogenesis (AHN) results in the generation of new neurons throughout adulthood, thereby contributing to key hippocampal functions such as learning and mood regulation. Here, we present a protocol for the immunofluorescence-based detection of AHN markers in fresh-frozen human hippocampal sections. We describe steps for tissue fixation, antibody incubation, and autofluorescence elimination. We then detail procedures for image acquisition and quantification. This protocol allows the reliable identification of distinct cell populations and could be adapted for broader immunohistochemical applications.

Before you begin

The mammalian hippocampus has the capacity to generate new neurons, namely dentate granule cells (DGCs), throughout life.^2,3 This process, known as adult hippocampal neurogenesis (AHN), is tightly regulated, confers enhanced neural plasticity, and is involved in key hippocampal functions such as learning and memory, and mood regulation.⁴ The occurrence of AHN has been reported in more than 120 mammalian species, including humans,^{3,5,6,7,8,9,10,11,12} yet a few studies failed to detect markers of neurogenesis in human adult hippocampi.^13,14,15 A putative source of discrepancy lies in the methodologies employed, as many studies rely on immunohistochemistry (IHC), a technique that is highly sensitive to tissue processing and preservation conditions. Recent work by our group showed that technical factors, including post-mortem delay and fixation time, critically affect the detection of AHN markers by IHC.^16,17 Additionally, the specificity and performance of antibodies continue to be a significant limitation, often leading to false negatives or ambiguous results, thus further complicating comparisons between studies. In this regard, the protocol outlined herein is intended to help establish consensus criteria that could contribute to increasing reproducibility across studies.

Here, we describe an optimized protocol that outlines the specific steps for the IHC detection of distinct cell markers related to AHN in fresh-frozen, glass-slide-mounted human hippocampal sections. This protocol was originally implemented to study how AHN is affected by neuropsychiatric disorders and distinct demographic and lifestyle-related factors in our related manuscript.¹ The cohort of subjects used in the aforementioned study corresponds to the neuropathology consortium of the Stanley Medical Research Institute (SMRI, USA) Brain Bank,^18,19 which includes 14 neurologically healthy control subjects, and 45 patients with major depression, bipolar disorder, and schizophrenia (15 per diagnostic group). High-quality tissue samples were obtained by medical examiners using a standardized procedure.¹⁹ One brain hemisphere was cut into 1.5 cm thick coronal sections, frozen in a mix of isopentane and dry ice, and stored at −70°C. The hippocampus was then embedded into Epredia M-1 Embedding Matrix and cut into 14-μm sections using a Leica cryostat. Chamber temperature was set to −17˚C, and specimen temperature was set to −14˚C. Subsequently, fresh-frozen sections were mounted on glass slides and stored at −80°C until use. Glass slides were previously coated using a mixture of 6g of porcine gelatin and 0.6g of Chromium(III) potassium sulfate dodecahydrate in 1200mls distilled water. Due to the size of the human hippocampal structure, 38 × 75 mm glass slides (larger than the standard 25 × 75 mm) were used.

We used the protocol described here to characterize the stages encompassed by AHN and also distinct populations that make up the neurogenic niche in human hippocampal samples (Figures 1, 2, and 3). Neural stem cells (NSCs) with radial-glia-like properties express vimentin and sex determining region Y-box 2 (Sox2) while lacking the mature astrocyte marker S100 calcium-binding protein β (S100β).^20,21 These cells are predominantly located in the subgranular zone (SGZ) and show a long vertical process that reaches the molecular layer (ML).^10,21 NSCs occasionally divide and give rise to transient amplifying progenitors and proliferative neuroblasts, which express doublecortin (DCX) and HuC-HuD and typically show an immature morphology with elongated nuclei and horizontal neurites.^10,22 As immature cells become committed to the neuronal lineage, they first express polysialylated neural cell adhesion molecule (PSA-NCAM) and, at later stages of maturation, calbindin (CB).^8,10,22 In the latter stages, they show a single, vertically oriented, primary apical dendrite and eventually become functionally integrated into hippocampal circuits.^8,10,22,23 To support the continuous generation of new neurons, the SGZ is composed of a specialized matrix, namely the neurogenic niche, characterized by a profuse vascular network and a high density of astrocytic and microglial cells. These cellular components establish complex crosstalk to regulate AHN. Mature astrocytes can be identified by the expression of S100β while microglial cells express ionized calcium-binding adaptor molecule 1 (Iba1).^10,24 Blood vessels and capillaries can be identified by the expression of Ulex Europaeus Agglutinin-I (UEA-1).^10,24 Beyond this application, this protocol can be easily adapted to process and analyze fresh-frozen human material for IHC studies.

Figure 1.

Experimental design

Briefly, (A) thaw and fix samples for 10 min in a fresh 4% formaldehyde solution. Subsequently, (B) draw a hydrophobic barrier surrounding the tissue to prevent solutions from spilling out, and (C) incubate the tissue with primary (24 h at 4°C) and secondary (2 h at rt (≈20°C–23°C)) antibodies. Afterwards, (D) stain cell nuclei using 4′,6-diamidino-2-phenylindole (DAPI) and (E) incubate the tissue with a commercial autofluorescence eliminator reagent to reduce autofluorescence. Next, (F) mount tissue sections and let them dry for at least 24 h before imaging. Finally, (G) acquire a determined number of stacks for each subject under a confocal microscope. Yellow scale bar: 500 μm. White scale bar: 20 μm.

Figure 2.

Image analysis

Overview of the stereological methods and morphometric determinations performed to quantify and characterize adult hippocampal neurogenesis in thin, fresh-frozen human samples. To obtain the reference volume (A–C), open confocal stacks in Fiji software (A), obtain z-projection images (B) and draw the area of interest using the freehand selection tool (C). Subsequently, cell densities are obtained (D–E) by counting the number of positive cells on each plane following stereological principles (D) and dividing the number obtained by the reference volume (E). Morphometric determinations (F–H) can be carried out on the same stacks acquired to estimate cell densities. The nuclear area of total dentate granule cells (DGCs) can be obtained by drawing the nuclear contour (DAPI channel) in the plane in which it is maximal, using the freehand selection tool (F). In doublecortin (DCX)⁺ immature neurons, the number and orientation of neurites can also be determined (G and H). To obtain the number, count the number of primary neurites that emerge from DCX⁺ cells (G). For the orientation, use the angle tool to measure the angle between the neurite and the hilar border (H). To analyze the dentate gyrus (DG) vasculature (I–K), obtain 40× immersion oil objective stacks (XY dimensions:159.72 μm, Z-interval: 1 μm). The thickness of UEA-1⁺ blood vessels (I(1)) can be determined by tracing a line perpendicular to the trajectory of the capillary. To determine the area occupied by UEA-1⁺ blood vessels (I(2)), apply an invariant threshold and measure the area above the selected threshold. Subsequently, divide the value obtained by the reference area (total (GCL+SGZ) or local (GCL or SGZ) (J). The vasculature skeleton (I(3)) can be semi-automatically traced using the Skeleton plug-in from Fiji to obtain the number of branches and junctions, the length, and the Euclidean distance (K). GCL: granule cell layer. SGZ: subgranular zone. Orange line delimits the area of interest (GCL and/or SGZ). White scale bar: 20 μm. Yellow triangle indicates a nucleus in the plane at which its size is maximal. Orange triangles indicate neurites in DCX⁺ cells. Red line indicates the hilar border of the GCL. Green triangles indicate UEA-1⁺ blood vessels. In I(1), yellow lines indicate the thickness of UEA-1⁺ blood vessels.

Figure 3.

Overview of expected outcomes

(A) Schematic representation of distinct cell populations related to adult hippocampal neurogenesis (AHN).

(B–H) Expected staining with anti- (B) vimentin, (C) sex determining region Y-box 2 (Sox2), (D) phospho-histone 3 (PH3), (E) HuC-HuD, (F) Doublecortin (DCX), (G) polysialylated-neural cell adhesion molecule (PSA-NCAM), and (H) calbindin (CB) antibodies.

(I) Schematic representation of the components of the human hippocampal neurogenic niche.

(J–M) Expected staining with anti- (J) S100 calcium-binding protein β (S100β), (K) ionized calcium-binding adaptor molecule 1 (Iba1), (L) phosphorylated γH2A.X (γH2A.X) antibodies, and (M) Ulex Europaeus Agglutinin-I (UEA-1). Yellow scale bar: 20 μm. White scale bar: 10 μm. Yellow triangles indicate positive cells. White triangles highlight the morphology of cells labelled for each marker.

Innovation

Unlike most exisiting studies which often rely on tissue samples fixed for long periods of time, this approach is specifically optimized for fresh-frozen human hippocampal samples. This adaptation prevents fixation-related epitope loss and preserves antigenicity across a broad range of neurogenic and niche-related markers (see section above). The protocol provides detailed, optimized instructions for the handling, storage, and immunohistochemical processing of human hippocampal sections. These refinements are intended to minimize technical variability and facilitate more reliable comparisons across samples and studies.

Institutional permissions

All experiments involving the use of human samples must be performed in accordance with relevant institutional and national guidelines and regulations. Samples were obtained following national laws and international ethical and technical guidelines on the use of human samples for biomedical research purposes. Brain tissue donation, processing, and use for research followed published protocols, which include approval by the Ethical Committees of the Instituto de Salud Carlos III (#CEI PI 30_2020# and #CEI PI 30_2020-V2-Ampliacion 2020#) and the Spanish Research Council (CSIC) (#025/2020# and #198/2020#).

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Rabbit monoclonal anti-doublecortin (EPR19997) (1:1000)	Abcam	Cat#ab207175; RRID: AB_2894710
Goat polyclonal anti-doublecortin (1:1000)	Santa Cruz	Cat#sc-8066; RRID: AB_2088494
Mouse monoclonal anti-polysialylated-neural cell adhesion molecule (PSA-NCAM) (1:1000)	Millipore	Cat#MAB5324; RRID:AB_95211
Rabbit polyclonal anti-phospho-histone 3 (PH3) (1:250)	Millipore	Cat#06-570; RRID:AB_310177
Rabbit polyclonal anti-calbindin D28k (CB) (1:2000)	Swant	Cat#CB38a; RRID: AB_2721225
Guinea pig polyclonal anti-S100 calcium-binding protein β (S100β) (1:500)	Synaptic Systems	Cat#287004; RRID:AB_2620025
Chicken polyclonal anti-vimentin (1:500)	Synaptic Systems	Cat#172006; RRID: AB_2800525
Goat polyclonal anti-SRY (sex-determining region Y)-box 2 (Sox2) (1:500)	R and D Systems	Cat# AF2018; RRID: AB_355110
Chicken monoclonal anti-ionized calcium-binding adaptor molecule 1 (Iba1) (1:500)	Synaptic Systems	Cat# 234 009; RRID: AB_2891282
Mouse monoclonal anti-phosphorylated γH2A.X (Ser139) (1:500)	Cell Signaling Technology	Cat#80312; RRID: AB_2799949
Mouse monoclonal anti-HuC-HuD (1:500)	Thermo Fisher Scientific	Cat#A21271; RRID: AB_221448
Alexa-488 anti-mouse (1:1000)	Thermo Fisher Scientific	Cat#A-21202; RRID: AB_141607
Alexa-488 anti-goat (1:1000)	Thermo Fisher Scientific	Cat#A-11055; RRID: AB_2534102
Alexa-488 anti-chicken (1:1000)	Thermo Fisher Scientific	Cat#A-11039; RRID: AB_2534096
Alexa-555 anti-mouse (1:1000)	Thermo Fisher Scientific	Cat#A-31570; RRID: AB_2536180
Alexa-555 anti-rabbit (1:1000)	Thermo Fisher Scientific	Cat#A-31572; RRID: AB_162543
Alexa-647 anti-mouse (1:1000)	Thermo Fisher Scientific	Cat#A-31571; RRID: AB_162542
Alexa-647 anti-rabbit (1:1000)	Thermo Fisher Scientific	Cat#A-31573; RRID: AB_2536183
Alexa-647 anti-goat (1:1000)	Thermo Fisher Scientific	Cat#A-21447; RRID: AB_2535864
Alexa-647 anti-guinea pig (1:1000)	Thermo Fisher Scientific	Cat#A-21450; RRID: AB_2735091
Biological samples
Human hippocampal sections obtained from 59 individuals (14 neurologically healthy control subjects, 15 patients with major depression (MD), 15 with schizophrenia (SCH), and 15 with bipolar disorder (BD).	Neuropathology consortium, Stanley Medical Research Institute (SMRI) Brain bank.	https://www.stanleyresearch.org/brain-research/neuropathology-consortium/
Chemicals, peptides, and recombinant proteins
Ulex Europaeus Agglutinin-1 (UEA-1) (1:750)	Vector Laboratories	Cat# B-1065; RRID: AB_2336766
Paraformaldehyde electron microscopy (EM) grade (16% wt/vol)	Electron Microscopy Sciences	Cat#15710
K2HPO4	Merck-Millipore	Cat#105101
NaH2PO4-H2O	Merck-Millipore	Cat#567545
Triton X-100	Sigma-Aldrich	Cat#93443
Bovine serum albumin (BSA)	Sigma-Aldrich	Cat#A7888
4′,6-diamidino-2-phenylindole (DAPI)	Merck-Millipore	Cat#268298
Ethanol (96%)	Merck-Millipore	Cat#159010
Mowiol 4-88	Merck-Millipore	Cat#475904
Glycerol (100%)	Sigma-Aldrich	Cat#G9012
Trizma base	Sigma-Aldrich	Cat#T1503
HCl	Merck-Millipore	Cat#113136
Epredia M-1 Embedding Matrix	Thermo Fisher Scientific	Cat#1310
Porcine gelatin	Sigma-Aldrich	Cat# G1890-100G
Chromium(III) potassium sulfate dodecahydrate	Sigma-Aldrich	243361-5G
Autofluorescence eliminator reagent	Merck-Millipore	Cat#2160
Software and algorithms
Image J 1.54i	Schneider et al.,25 and Schindelin et al;26	https://imagej.nih.gov/ij/
DG vasculature analysis (Fiji Macro)	Rust et al.,27	Supplementary material of Rust et al.,27
Other
ImmEdge Hydrophobic Barrier PAP Pen	Vector Laboratories	Cat#H4000
Glass staining dish	BRAND	Cat#472200
Glass tray	BRAND	Cat#472000
40 × 50 mm microscopy glass coverslips	Thermo Fisher Scientific	Cat#BC0400050A140MNZ0
LSM 900 Zeiss confocal microscope	Carl Zeiss	https://www.zeiss.com/microscopy/es/home.html
Immersol™ 518 F (Immersion oil)	Carl Zeiss	Cat#444960-0000-000
Vectashield mounting medium	Vector	H-1000-10

Materials and equipment

0.2 N phosphate buffer

To prepare 1 L of this solution, add 28 g of K₂HPO₄ to 800 mL of bidistilled water and stir until dissolved completely. Subsequently, add 5.3 g of NaH₂PO₄-H₂O and stir. Adjust pH to 7.4 and bring the final volume to 1 L with bidistilled water.

Reagent	Final concentration	Amount
K2HPO4	2.8% (w/v)	28 g
NaH2PO4-H2O	0.53% (w/v)	5.3 g
ddH2O	N/A	Up to 1 L
Total	N/A	1 L

Store at room temperature (rt) (≈20°C–23°C) for up to 6 months.

0.1 N PB

Dilute the 0.2 N PB solution in bidistilled water at a 1:1 vol/vol ratio. This solution can be stored at rt (≈20°C–23°C) for up to 6 months.

4% formaldehyde fixative solution

Dilute the commercial 16% formaldehyde solution in 0.2 N PB and distilled water at a 1:2:1 vol/vol/vol ratio. Adjust pH to 7.4 if necessary.

Reagent	Final concentration	Amount
16% formaldehyde	4%	25 mL
0.2 N PB	N/A	50 mL
dH2O	N/A	25 mL
Total	N/A	100 mL

Prepare this solution immediately before use.

CRITICAL: Formaldehyde is toxic, mutagenic, carcinogenic, and irritant. Perform all procedures involving this solution under a chemical fume hood and wear appropriate personal protective equipment (PPE), including gloves, mask, and goggles. Dispose of the formaldehyde waste according to your institution guidelines.

PB-Triton X-100-bovine serum albumin 0.5% solution

Mix all components on a magnetic shaker until completely dissolved.

Reagent	Final concentration	Amount
BSA	0.5% (w/v)	1 g
Triton-X-100	0.5% (v/v)	1 mL
0.1 N PB	N/A	Up to 200 mL
Total	N/A	200 mL

Store at 4°C for up to 1 week.

CRITICAL: Triton-X-100 is an irritant. Wear appropriate PPE, including gloves, a mask, and goggles, when performing procedures involving the manipulation of this reagent.

70% ethanol solution

To prepare 1 L of this solution, mix 729 mL of 96% ethanol with 271 mL of bidistilled water. This solution can be stored at rt (≈20°C–23°C) for up to 3 months.

CRITICAL: Ethanol (96%) is an irritant. Wear appropriate PPE, including gloves, a mask, and goggles, when performing procedures involving this alcohol.

0.2 M Tris-HCl (pH = 8.5)

To prepare 500 mL of this solution, add 12.114 g of Trizma base to 350 mL of bidistilled water. Adjust pH to 8.5 by adding HCl. Add bidistilled water up to a volume of 500 mL. This solution can be stored at rt (≈20°C–23°C) for up to 6 months.

CRITICAL: HCl is toxic if swallowed. Perform all procedures involving the manipulation of this reagent under a chemical fume hood and wear appropriate PPE, including gloves, mask, and goggles.

Antifade mounting medium

To prepare ≈18 mL of non-commercial fluorescence anti-fade mounting medium.

• •Add 6 g of 100% glycerol to 6 mL of distilled water and mix.
• •Subsequently, add 2.4 g of Mowiol 4-88 and mix overnight (≈16 h) in an orbital shaker at rt (≈20°C–23°C).
• •Add 12 mL of 0.2 M Tris-HCl (pH = 8.5) and mix for 10 min in an orbital shaker at 50°C.
• •Centrifuge at 5000g for 12 min at 4°C and aliquot the supernatant into 2-mL Eppendorf tubes.

Reagent	Final concentration	Amount
100% Glycerol	33.3% (w/v)	6 g
Mowiol 4–88	13.3% (w/v)	2.4 g
dH2O	N/A	6 mL
Tris-HCl 0.2 M (pH = 8.5)	N/A	12 mL
Total	N/A	18 mL

Store aliquots at −20°C for up to 1 year.

CRITICAL: Glycerol is an irritant, and HCl is toxic if swallowed. Wear appropriate PPE, including gloves, a mask, and goggles, when performing procedures involving the manipulation of these reagents.

Alternatives: A commercial mounting medium such as Vectashield can also be used, provided it is suitable for the microscope and objectives being used.

Step-by-step method details

Tissue fixation

Timing: 30 min

A short fixation step ensures structural integrity of the tissue, while maintaining epitope accessibility to antibodies (Figure 1A). Fixation time should be optimized according to the target epitope, as some may require longer or shorter fixation.

• 1.Thaw fresh-frozen glass-slide-mounted sections at rt (≈20°C–23°C) for 10 min.
• 2.
  Fix sections in a freshly prepared 4% formaldehyde solution.

  • a.
    Place slides in a clean glass tray.
  • b.
    Fill a glass staining dish with 4% formaldehyde solution and immerse the tray containing the slides for 10 min.

CRITICAL: Formaldehyde is toxic, mutagenic, carcinogenic, and irritant. Perform all procedures involving this solution under a chemical fume hood and wear appropriate personal protective equipment (PPE), including gloves, mask, and goggles. Dispose of the formaldehyde waste according to your institution guidelines.

• 3.Wash the sections three times for 3 min each in 0.1 N PB solution. For each of the washes, fill a new glass staining dish with 0.1 N PB solution and carefully transfer the tray containing the slides to the new dish.

Primary and secondary antibody incubation

Timing: 28 h

The following steps are performed to permeabilize, block, and incubate tissue sections with primary and secondary antibodies to detect specific cell populations (Figures 1B and 1C).

• 4.Dry the section using a small piece of paper towel.

CRITICAL: Take extreme care not to damage the tissue. First, dry the side of the glass slide that does not contain the tissue to remove most of the excess liquid. Then, without touching the tissue itself, gently remove the remaining solution from the side containing the tissue by carefully pressing the paper towel around its perimeter. From this step onwards, repeat these steps every time drying is required. Sections should not be left to completely dry out.

• 5.Using the ImmEdge Hydrophobic Barrier PAP Pen, draw an area surrounding each section to ensure that the distinct solutions do not spill during incubations (see troubleshooting – problem 1).
• 6.Prepare a humidity chamber by adding a wet paper towel to the bottom of an incubation box. This will prevent tissue from drying and solutions from evaporating during long incubations.
• 7.Place tissue sections in the humidity chamber with the tissue facing upwards. At this point, place 1 mL of 0.1 N PB onto each section to prevent the tissue from drying during the next step.
• 8.Prepare the primary antibody solution by diluting primary antibodies in the incubation buffer (0.5% PB-T-BSA), which contains 0.5% Triton X-100 and 0.5% BSA diluted in 0.1 N PB. Dilute the primary antibodies at the concentration recommended by the manufacturer.
• 9.Decant the previous solution and dry the section using a small piece of paper towel (see step 4 for specific instructions).
• 10.Add 300 μL of primary antibody solution. Adapt the volume needed to the specific size of the tissue.

CRITICAL: The indicated volume is optimized for tissue sections measuring approximately 50 × 35 mm. To prevent uneven staining, ensure that the primary antibody solution fully covers the entire tissue section and that no bubbles are formed. Add solutions to the tissue gently to prevent sections from detaching from the slides (see troubleshooting—problem 2).

• 11.Place the lid on the humidity chamber and incubate primary antibodies for 24 h at 4°C (see troubleshooting—problem 3).
• 12.Remove primary antibody solution by decantation.
• 13.Place slides in a clean glass tray.
• 14.Fill a glass staining dish with 0.5% PB-T-BSA solution and carefully transfer the tray containing the slides to the new dish. Wash slides for 3 min.
• 15.Take slides out of the glass tray and transfer them to the humidity chamber.
• 16.Gently wash tissue sections twice for 3 min each by adding 1 mL of 0.5% PB-T-BSA.

CRITICAL: Do not decant the liquid of the last wash until the next buffer is prepared, to avoid tissue drying.

• 17.Prepare the secondary antibody solution by diluting the secondary antibodies in 0.5% PB-T-BSA incubation buffer. This protocol is optimized for the use of Alexa®-conjugated secondary antibodies (see key resources table). Dilute secondary antibodies at the concentration recommended by the manufacturer.
• 18.Decant the previous solution and dry the section using a small piece of paper towel (see step 4 for specific instructions).
• 19.Add 500 μl of secondary antibody solution. Adapt the volume needed to the specific size of the tissue.

CRITICAL: The indicated volume is optimized for tissue sections measuring approximately 50 × 35 mm. To prevent uneven staining, ensure that the secondary antibody solution fully covers the entire tissue section and that no bubbles are formed.

• 20.Place the lid on the humidity chamber and incubate the secondary antibodies for 2 h at rt (≈20°C–23°C).

Note: Some secondary antibodies are light-sensitive. Therefore, from this step on, protect the humidity chamber from light. To this end, aluminum foil can be used to cover the chamber.

• 21.
  Gently wash tissue sections three times for 3 min each by adding 1 mL of 0.5% PB-T-BSA.

  CRITICAL: Do not decant the liquid of the last wash until the next buffer is prepared, to avoid tissue drying.

  Note: Appropiate controls should be included in every immunostaining experiment to confirm antibody specificity and identify non-specific staining. These may include:

  • a.
    Positive control: when possible, include a tissue known to express the protein of interest.
  • b.
    Negative control: when possible, include a tissue known to not express the protein of interest.
  • c.
    No-primary antibody control: incubate the section with the primary antibody diluent alone, followed by the secondary antibody and detection reagents. This control confirms that the observed signal originates from the primary antibody binding to its antigen.
  • d.
    Secondary antibody control: in multichannel experiments, apply each secondary antibody individually to ensure it does not recognize other primary antibodies, bind to other secondary antibodies, or generate bleed-through between channels.
  • e.
    Pre-absortion control: pre-absorb the primary antibody with a synthetic peptide corresponding to the immunogen before incubation. Sections are then processed with the pre-absorbed primary antibody, followed by the secondary antibody and detection reagents. When the antibody is specific, little to no staining should be observed in this control.
    Refer to troubleshooting—problems 4, 5, 6, and 7 if staining issues are encountered.

Nuclear staining

Timing: 15 min

Complete steps 22–25 to stain nuclei (Figure 1D).

• 22.Dilute 4′,6-diamidino-2-phenylindole (DAPI) at a 1 μg/mL concentration in 0.1 N PB.

CRITICAL: Chemical dyes with DNA-binding capacity are mutagenic and toxic. DAPI should be handled as a potential mutagen. Wear appropriate PPE, including gloves and a mask, when handling this dye. Discard all waste in appropriate containers.

• 23.Decant the previous solution and dry the section using a small piece of paper towel (see step 4 for specific instructions).
• 24.Gently add 500 μl of the DAPI solution and incubate for 10 min. Adapt the volume needed to the specific size of the tissue.
• 25.Rinse the tissue sections three times by adding 1 mL of 0.1 N PB and incubating for 3 min.

Autofluorescence elimination

Timing: 15 min

Incubation with a commercial autofluorescence eliminator reagent allows a significant reduction of background noise, thereby improving the signal-to-noise ratio (Figure 1E).

• 26.Dry the section using a small piece of paper towel (see step 4 for specific instructions).
• 27.Gently add 1 mL of a 70% ethanol solution over the tissue and incubate for 2 min at rt (≈20°C–23°C).
• 28.Remove the ethanol solution by decantation.
• 29.Gently add 1 mL of the commercial autofluorescence eliminator reagent (see key resources table) over the tissue and incubate for 2 min at rt (≈20°C–23°C) (see troubleshooting — problem 8).

CRITICAL: Autofluorescence eliminator reagent is toxic, carcinogenic, irritant, and flammable. Perform all procedures under a chemical fume hood when handling this reagent and wear appropriate PPE, including gloves, mask, and goggles.

• 30.Remove the reagent by decantation.
• 31.Wash the sections three times for 1 min each with 70% ethanol solution by gently adding 1 mL of solution over the tissue. Remove washing solution by decanting the solution.
• 32.Wash the sections three times for 1 min each with a 0.1 N PB solution by gently adding 1 mL of solution over the tissue. Remove washing solution by decanting the solution.
• 33.Place slides on a glass tray and immerse the tray in a glass staining dish filled with 0.1 N PB solution. Cover the glass staining dish with aluminum foil to protect it from light until the section is removed for mounting (see the next step).

Mounting

Timing: 5 min (per slide) and 24 h to dry

Complete steps 34–39 to mount stained sections (Figure 1F).

• 34.Take one slide out of the glass staining dish.
• 35.Dry the section using a small piece of paper towel by going around the tissue limits (see step 4 for specific instructions).
• 36.Take a coverslip and distribute approximately 300 μL of mounting medium on one of the sides. Use of a glass rod is recommended for an even distribution of the mounting medium.

Note: Adapt the volume needed to the specific size of the tissue.

• 37.With the help of fine curved-tip tweezers, gently place the coverslip on top of the tissue, with the side that contains the mounting medium facing the tissue.
• 38.Once mounted, gently press the coverslip to avoid trapping air bubbles and to ensure an even distribution of mounting medium.
• 39.Place the slide on a dry chamber protected from light and leave to dry at rt (≈20°C–23°C) for at least 24 h before imaging.

Note: Repeat steps 34–39 for each tissue slide.

Confocal image acquisition

Timing: Variable

Following IHC, confocal image acquisition should be performed immediately, as fluorescence fading may occur over time. In that regard, confocal images should ideally be acquired within two weeks after tissue mounting. In the related manuscript,¹ a LSM 900 Zeiss confocal microscope (Carl Zeiss) equipped with Zen Blue software (version 3.10.103.04000) was used for imaging purposes. When acquiring confocal images, researchers should ideally be blinded to experimental conditions (Figure 1G).

• 40.Place the glass slide on the confocal microscope.
• 41.Use a low-magnification dry objective and the epifluorescence function to locate the dentate gyrus (DG) using the DAPI channel.

Note: Select the lowest fluorescence intensity on the lamp to avoid fluorescence quenching.

• 42.Once the DG has been identified, use a Plan-Apochromat 40X1.3 Oil DIC (UV) VIS IR-M27 objective to focus on the granule cell layer (GCL) and use the DAPI channel to select the sampling zone.
• 43.Adjust laser power, gain, and acquisition setting parameters for each channel.
• 44.Set the dimensions of the sampling zone and the z-step.

Note: These parameters should be optimally adjusted depending on the microscope and objective used, as well as on the parameters/cell populations of interest. In the related manuscript,¹ to estimate the density of total DGCs, high-power magnification stacks were obtained under a Plan-Apochromat 63× / 1.40 Oil DIC M27 objective with a 2× zoom (1024×1024 pixels; XY dimensions: 50.7 μm; pixel size: 49.5 nm; Z-step: 1 μm; pinhole dimensions: 0.9 Airy units). To estimate the density of Sox2⁺, PH3⁺, vimentin⁺, DCX⁺, HuC-HuD⁺, PSA-NCAM⁺, CB⁺, S100β⁺, Iba1⁺, and γH2AX⁺ cells, as well as to study UEA-1⁺ blood vessels, stacks were obtained under a Plan-Apochromat 40×/1.3 Oil DIC (UV) VIS IR-M27 objective with a 0.7 – 1× zoom (1024×1024 pixels; XY dimensions: 234.89–159.72 μm; pixel size: 229.5–155.9 nm; Z-step: 1 μm; pinhole dimensions: 0.99 Airy units). Fluorophores were excited using 405, 488, 561 and 640 nm laser lines. Emssion was collected at 456 nm (DAPI), 488–517 nm (Alexa-488), 555–568 nm (Alexa-555), and 647–668 nm (Alexa-647), respectively. Fluorescence signals were detected using GaAsP photomultiplier tube (PMT) detectors. Laser power was adjusted for each fluorophore and staining to optimize signal-to-noise ratio while avoiding saturation and photobleaching.

• 45.Define the z-stack thickness. In the related manuscript,¹ the stacks contained the full volume of the tissue.
• 46.Acquire a variable number of stacks of images per subject.

Note: This number should be adjusted to the relative abundance and specific characteristics of the cells/structures of interest. Sampling should be randomly distributed along the DG. In the related manuscript,¹ 10 stacks of images per subject were acquired to estimate cell densities and perform morphometric determinations.

Note: Refer to troubleshooting — problem 9 if issues with image acquisition are encountered.

Image analysis

Timing: Variable

This section describes the necessary steps to estimate total and local cell densities, as well as distinct morphometric characteristics of various AHN-related cell populations. To this end, the free software Fiji (Fiji Is Just Image J)^25,26 was used in the related manuscript¹ as further described below (Figure 2).

• 47.
  Determine the reference volume (Figures 2A–2C).

  • a.
    Obtain a maximum z-projection image (Image > Stacks > Z Project; Projection type: Max intensity) for each stack of images (Figure 2B).
  • b.
    In the z-projection image, trace the region of interest in the DAPI channel using the freehand selection tool from Fiji.

    Note: Tracing should encompass the area in which positive cells will be counted. To estimate total cell densities, delineate the area of the GCL plus the SGZ. To determine local cell densities, trace the area of the GCL and SGZ separately (Figure 2C).

  • c.
    Measure the area and multiply the obtained value by the z-thickness of the stack to determine the reference volume.

    Note: To estimate the density of total DGCs, use high-power magnification images including a small portion of the GCL, and calculate the reference volume by multiplying the X, Y, and Z dimensions of the stack.

• 48.
  Use the physical dissector method adapted to confocal microscopy to estimate cell densities²⁸ (Figures 2D and 2E).

  • a.
    Open the confocal stack of images in Fiji.
  • b.
    Select the “Cell counter” tool (Plugins > Analyze > Cell Counter) and click on “Initialize”.
  • c.
    Count the number of positive cells that are inside the reference volume on individual planes. Count positive cells in the first plane where they appear and avoid counting the same cell in the following planes. To avoid overestimating the number of cells, three inclusion faces and three exclusion faces of the stack (X, Y, and Z dimensions) should be defined. Cells that touch inclusion faces (i.e., left, back, and bottom) should be included in the count, whereas those that touch exclusion faces (i.e., right, front, and top) should be excluded. Cell counts can be saved using the option “Save Markers” and reopened whenever needed (Figure 2D).

    CRITICAL: Prior to this step, clear and objective criteria to unambiguously identify positive cells must be defined. These criteria can be based on previously published and validated data. In cases where this information is not available, such as when identifying novel markers, the following criteria should be taken into account. Cell morphology must be unequivocal, and any cells displaying apoptotic features on the DAPI channel or autofluorescence in any of the acquired channels should be excluded from the count. Apoptotic nuclei may appear smaller and more intensely stained than healthy nuclei due to chromatin condensation. They typically exhibit fragmented or irregular shapes, with chromatin appearing as condensed clumps.

  • d.
    Divide the number of positive cells by the reference volume to obtain cell densities (number of cells/mm³) (Figure 2E).
• 49.
  Morphometric determinations. Distinct parameters such as the nuclear area, number, and orientation of neurites can be determined using the same images acquired for cell density estimation (Figures 2F–2H).

  • a.
    To estimate the nuclear area of DGCs, use high-power magnification stacks obtained under a Plan-Apochromat 63×/1.40 Oil DIC M27 objective with a 2× zoom (1024×1024 pixels; XY dimensions: 50.7 μm; pixel size: 49.5 nm; Z-step: 1 μm; pinhole dimensions: 0.9 Airy units). In the DAPI channel, draw the nuclear contour in the plane in which it is maximal using the freehand selection tool from Fiji (Figure 2F). Measure at least 50 cells per subject.
  • b.
    To determine the number of neurites in DCX⁺ cells, count the number of primary neurites that emerge directly from the soma (Figure 2G).
  • c.
    Estimate the orientation of neurites in DCX⁺ cells by determining the angle formed by the hilar border of the GCL and the neurite (Figure 2H).

Note: A neurite is considered to have a horizontal orientation (parallel to the SGZ) if the angle is ≤45°, whereas it is considered vertical when the angle is between 45° and 90°.

• 50.
  DG vasculature analysis. Distinct parameters of UEA-1⁺ blood vessels can be determined using a Plan-Apochromat 40×/1.3 Oil DIC (UV) VIS IR-M27 objective (1024×1024 pixels; XY dimensions: 159.72 μm; pixel size: 155.9 nm; Z-step: 1 μm; pinhole dimensions: 0.99 Airy units (Figures 2I–2K).

  • a.
    To determine the thickness of UEA-1⁺ blood vessels, trace and measure a line perpendicular to the trajectory of the capillary using the straight-line tool from Fiji. Measure at least 100 capillaries per subject. If needed, digitally increase the magnification of the confocal stack to facilitate measurement (Figure 2I(1)).
  • b.
    To determine the area of the DG occupied by UEA-1⁺ blood vessels, obtain at least 10 stacks of confocal images per subject. All the stacks must contain the same number of planes.

    • i.
      Obtain a maximum z-projection image (see step 47a) for each stack.
    • ii.
      Apply an invariant threshold to z-projection images (Image > Adjust > Threshold) (Figure 2I(2)).
    • iii.
      Next, using the DAPI channel, trace and measure either total (SGZ+GCL) or local (SGZ or GCL) areas (reference area).
    • iv.
      Subsequently, measure the area above threshold within the local or total area.
    • v.
      Divide the value obtained by the reference area to determine the area occupied by capillaries (Figure 2J).
  • c.
    To analyze the vascular skeleton, a semi-automated method in Fiji adapted from Rust et al.²¹ can be used. Full details on the application and the corresponding Fiji macros are available on the referenced publication (see key resources table).

    • i.
      Briefly, regions of interest (ROIs) of a determined and invariable size are selected from each confocal stack.

      Note: In the related manuscript,¹ 90 × 28 μm ROIs were used.

    • ii.
      Then, the Fiji macro applies an adjustable threshold, after which small artifacts are removed before image binarization.
    • iii.
      Subsequently, the Skeleton plugin from Fiji can be used to automatically trace the vascular skeleton (Figure 2I(3)) and extract several parameters such as length, number of branches, and junctions or Euclidean distance (length of the line segment between the beginning and the end of the branch) (Figure 2K). The tortuosity index can be obtained by dividing the branch length by the Euclidean distance, minus 1 to account for the difference.

Expected outcomes

The use of this protocol allows the stereological estimation of cell densities related to AHN and the neurogenic niche, as well as morphometric determinations and an in-depth analysis of the DG vasculature on post-mortem human hippocampal samples. Both single and double/triple immunostaining can be performed, thereby maximizing the use of human tissue. Representative staining patterns for the labelling of each distinct cell population are shown in Figure 3. Some of the expected outcomes include clear identification of Vimentin⁺ S100β^- or Sox2⁺ NSCs (Figures 3B and 3C). Co-labelling of Vimentin⁺ S100β^- cells with PH3 (Figure 3D) allows the identification of the proliferating NSC pool in the adult human dentate gyrus. Proliferating neuroblasts can be identified by the expression of HuC-HuD (Figure 3E), whereas immature neurons at distinct maturation stages can be identified by the expression of DCX and PSA-NCAM (Figures 3F and 3G). Mature DGCs can be reliably identified by the expression of CB (Figure 3H). Furthermore, this method allows the identification of key components of the neurogenic niche, such as S100β⁺ astrocytes (Figure 3J), Iba1⁺ microglia (Figure 3K), and UEA-1⁺ blood vessels (Figure 3M), as well as cells positive for γH2A.X (Figure 3L), a marker of DNA damage/repair.

In summary, this protocol offers a comprehensive toolbox to study AHN in fresh-frozen post-mortem human tissue and is expected to contribute to broadening our understanding of AHN in physiological and pathological conditions.

Limitations

The protocol presented here allows high-resolution immunofluorescence of fresh-frozen, thin hippocampal slices. However, several potential limitations must be taken into account. First, conventional procedures used at most brain banks worldwide typically include extended tissue fixation in formalin, which can impair the detection of AHN markers by IHC.^16,17 Hence, the availability of high-quality, well-preserved human brain samples is a potential major limitation. Moreover, all studies involving the use of human samples should be performed following national and international guidelines (see institutional permissions), and, therefore, the time required to obtain such authorization should be considered when designing experiments.

In addition, a general limitation of research based on post-mortem human material is inter-individual variability (pertaining, e.g., to lifestyle, habits, comorbidities, ante-mortem conditions, among others). In this regard, the use of large cohorts of subjects is highly recommended to minimize the impact of the aforementioned factors. Moreover, reporting all available epidemiological factors is strongly recommended to enable data comparison across different studies and laboratories.

This protocol has been validated to be used with a wide variety of antibodies related to AHN (see key resources table). While it can be applied to detect other cell populations, proper antibody validation is essential to ensure signal specificity and reproducibility. Researchers should have demonstrated proficiency with IHC techniques to ensure the identification of signal specificity vs. background or autofluorescence, for instance. Moreover, exhaustive antibody validation is recommended and might include testing multiple antibodies to detect the same target. Therefore, considerable costs and time dedication of researchers should be anticipated.

Troubleshooting

Problem 1

Solutions are running outside the tissue barrier. (Refer to step 5).

Potential solution

Re-draw the hydrophobic barrier using the PAP pen to contain solutions. Redrawing can be performed at any time when needed during the protocol.

Problem 2

Tissue is detached from glass slides. (Refer to step 10).

Potential solution

Take extra care to gently add reagents or solutions. Using a pipette, let drops fall on top of the tissue without applying too much pressure and avoid hitting tissue edges to prevent mechanical disruption. In addition, coated slides, either commercial or custom-made (i.e., 2% gelatin-coated), can be used to improve tissue adhesion.

Problem 3

Tissue is dry after incubation. (Refer to step 11).

Potential solution

Ensure the humidity chamber is being used properly. Additional wet paper can be placed inside the chamber to maintain humidity. Moreover, ensure that a sufficient volume of solution is placed on the tissue.

Problem 4

Lack of signal when using previously validated antibodies. (Refer to steps 4–21).

Potential solution

The antibody may have lost efficiency over time. Test it on a positive control tissue and consider adding and/or optimizing an antigen retrieval step to improve epitope exposure to the primary antibody.

Problem 5

Lack of signal when using previously unvalidated antibodies. (Refer to steps 4–21).

Potential solution

The antibody may not work in human tissue. Check the manufacturer's website to determine whether the antibody is designed against the human epitope. Test it on a human positive control tissue sample if available, and/or consider adding an antigen retrieval step to improve epitope exposure to the primary antibody.

Problem 6

Staining has high background but presence of specific signal. (Refer to steps 4–21).

Potential solution

Concentration of the antibody might be too high. Test lower concentrations until an acceptablen background/signal ratio is achieved. Consider including a blocking step to prevent non-specific binding of antibodies. Alternatively, incubation time might be longer than needed.

Problem 7

Staining shows specific signal, but with low intensity. (Refer to steps 4–21).

Potential solution

Concentration of the antibody might be too low. Test higher concentrations until an acceptable background/signal ratio is achieved. Alternatively, incubation time might be shorter than needed.

Problem 8

Staining shows presence of autofluorescence. (Refer to step 29).

Potential solution

Increase incubation time with the autofluorescence eliminator reagent.

Problem 9

Fluorescence quenching. This can be identified by the observation of a progressive loss of fluorescence intensity over time. For example, a previously bright structure may appear faded or even disappear after repeated imaging. (Refer to steps 40–46).

Potential solution

If fluorescence quenching occurs during sample observation under the microscope, attempt lower exposure of the sample to the fluorescent light. To this end, reduce the intensity of the fluorescent lamp and minimize observation time.

If fluorescence quenching occurs during imaging, acquisition parameters may need to be optimized. These adjustments include, but are not limited to, reducing laser power and increasing gain instead, reducing line averaging, or reducing image resolution.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, María Llorens-Martín (m.llorens@csic.es).

Technical contact

Technical questions on executing this protocol should be directed to and will be answered by the technical contact, María Llorens-Martín (m.llorens@csic.es).

Materials availability

No original materials have been generated in this study. Requests for human brain samples should be addressed to the Stanley Medical Research Institute (stanleyresearch.org).

Data and code availability

• •This study did not generate datasets.
• •This paper does not report original code.

Acknowledgments

The authors would like to thank the patients and their families for generously donating brain samples. Moreover, they wish to thank the SMRI brain bank for providing samples and M. Flor-García, J. Terreros-Roncal, E. Moreno-Jiménez, C. Rodríguez-Moreno, and the Confocal Microscopy Facility at the CBMSO for technical assistance.

This study was supported by the following funding sources: the European Research Council (ERC) (ERC-CoG-2020-101001916) (M.L.-M.), the Spanish Ministry of Economy and Competitiveness (PID2020-113007RB-I00 and PID2023-146572OB-I00) (M.L.-M.), the BrightFocus Foundation (A2024021S) (M.L.-M.), and the Center for Networked Biomedical Research on Neurodegenerative Diseases (CIBERNED, Spain) (M.L.-M.). The salary of B.M.-V. was supported by postdoctoral fellowships awarded by the Consejo Nacional de Ciencia y Tecnología (CONACYT) of the Mexican Government (385084) and the Secretaria de Educación, Ciencia Tecnología e Innovación (SECTEI) of the Regional Government of Ciudad de México (CDMX) (CM-SECTEI/159/2021). The salary of M.G.-C. was supported by a Formación de Personal Investigador (FPI) contract, associated with the PID2020-113007RB-I00 grant (M.L.-M.), awarded by the Spanish Ministry of Economy and Competitiveness (PRE2021-097690).

Author contributions

B.M.-V., M.G.-C., and M.L.-M. conceived and designed the study. B.M.-V. and M.G.-C. performed the experiments. The Stanley Medical Research Institute provided the brain samples. B.M.-V., M.G.-C., and M.L.-M. analyzed the data. M.G.-C. and M.L.-M. designed the figures and wrote the initial manuscript. B.M.-V. and M.L.-M. obtained funding. All the authors discussed the data and revised the final version of the manuscript.

Declaration of interests

The authors declare no competing interests.