How do different tissue culture matrices affect background in fluorescent microscopy?

How do different tissue culture matrices affect background in fluorescent microscopy?

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In response to my previous question, I've been reading up a little bit on poly-D-lysine, Collagen I, Collagen IV, laminin, and other tissue culture coatings that promote cell adhesion. I've always assumed that anything other than standard TC-treated plastic or glass would significantly increase background, but perhaps my views on background fluorescence are a little outdated. Does anybody have experience with these in a fluorescent microscopy/high-throughput screening environment?

Specifically in my case, I'm looking at endocytosis and trafficking of a labeled protein into the lysosome. I'm labeling the protein with the pH-dependent dye pHrodoTM from Molecular Probes, which supposedly has very little fluorescence at neutral pH, but becomes very bright as the pH drops when endocytic vesicles become lysosomes. This theoretically means that a final wash step is not needed, but with a matrix coating on the plates I'm worried about background.

So, what is the current thinking as far as background fluorescence of the various TC matrices is concerned? Does the background come from the matrix itself, or by the fluorescent dye becoming adsorbed to it? Is it wavelength-dependent? Fortunately I may not be stuck with my poorly-adhering cells, and I may not need supplemental matrix at all in the end, but I still want a better understanding of how it works.

Extracellular matrix (ECM) fluoresces, especially Collagen and Laminin. The maximum is in the DAPI and FITC channels and the fluorescence becomes weaker towards longer wavelengths. However, since the coat on the TC flasks is very thin, I would not expect this to be a problem. The best thing is just to try it. There is also a quite famous document available which might be of help:

Autofluorescence: Causes and Cures

Multispectral imaging in biology and medicine: Slices of life †

Multispectral imaging (MSI) is currently in a period of transition from its role as an exotic technique to its being offered in one form or another by all the major microscopy manufacturers. This is because it provides solutions to some of the major challenges in fluorescence-based imaging, namely ameliorating the consequences of the presence of autofluorescence and the need to easily accommodate relatively high levels of signal multiplexing. MSI, which spectrally characterizes and computationally eliminates autofluorescence, enhances the signal-to-background dramatically, revealing otherwise obscured targets. While this article concentrates on examples derived from liquid-crystal tunable filter-based technology, the intent is to showcase the advantages of multispectral imaging in general. Some technologies used to generate multispectral images are compatible with only particular optical configurations, such as point-scanning laser confocal microscopy. Band-sequential approaches, such as those afforded by liquid-crystal tunable filters (LCTFs), can be conveniently coupled with a variety of imaging modalities, which, in addition to fluorescence microscopy, include brightfield (nonfluorescent) microscopy as well as small-animal, noninvasive in-vivo imaging. Brightfield microscopy is the chosen format for histopathology, which relies on immunohistochemistry to provide molecularly resolved clinical information. However, in contrast to fluorescent labels, multiple chromogens, if they spatially overlap, are much harder to separate and quantitate, unless MSI approaches are used. In-vivo imaging is a rapidly growing field with applications in basic biology, drug discovery, and clinical medicine. The sensitivity of fluorescence-based in-vivo imaging, as with fluorescence microscopy, can be limited by the presence of significant autofluorescence, a limitation which can be overcome through the utilization of MSI. © 2006 International Society for Analytical Cytology

Multispectral imaging is the acquisition of spectrally resolved information at each pixel of an imaged scene. Many different technologies can be employed to generate such information, ranging from multi-position filter-wheels, gratings and prisms, laser-scanning single point spectrographs, electronically adjustable tunable filters, Fourier-transform imaging spectrometry, and computed tomographic imaging spectroscopy (reviewed in Ref. ( 1 )). This report will highlight the use of liquid crystal tunable filter- (LCTF-) based multispectral imaging approaches, along with application-specific analysis tools, for a variety of imaging tasks, but should also be read as a presentation of the advantages of MSI in general.

REVIEW article

  • Department of Plastic and Hand Surgery, University Hospital of Erlangen, Friedrich𠄺lexander University Erlangen–Nürnberg (FAU), Erlangen, Germany

Intravital microscopy (IVM) study approach offers several advantages over in vitro, ex vivo, and 3D models. IVM provides real-time imaging of cellular events, which provides us a comprehensive picture of dynamic processes. Rapid improvement in microscopy techniques has permitted deep tissue imaging at a higher resolution. Advances in fluorescence tagging methods enable tracking of specific cell types. Moreover, IVM can serve as an important tool to study different stages of tissue regeneration processes. Furthermore, the compatibility of different tissue engineered constructs can be analyzed. IVM is also a promising approach to investigate host reactions on implanted biomaterials. IVM can provide instant feedback for improvising tissue engineering strategies. In this review, we aim to provide an overview of the requirements and applications of different IVM approaches. First, we will discuss the history of IVM development, and then we will provide an overview of available optical modalities including the pros and cons. Later, we will summarize different fluorescence labeling methods. In the final section, we will discuss well-established chronic and acute IVM models for different organs.

Fig. 2

The absolute fluorescence intensities differ significantly between normal ( n = 36 ), perilesional ( n = 30 ), and cancerous ( n = 52 ) lung tissue. The significance is presented by “*” ( p < 0.05 ), “**” ( p < 0.01 ), and “***” ( p < 0.001 ).

Tissue clearing and GFP preservation in the heart

The fast development and usage of tissue clearing techniques relates to the rapid development of imaging methods, such as confocal microscopy and light sheet microscopy. Together these methods can be used to reconstruct the 3D anatomy of the tissue (Costantini etਊl., 2019).

One of the main model organisms in biological research is the mouse model, with its wide array of possible genetical modification including insertion of fluorescently labeled proteins (e.g., GFP) as reporter genes. The first developed tissue clearing methods used hydrophobic compounds, which dehydrate the tissue and dehydration quench the introduced fluorescent proteins (i.e., GFP). To avoid this phenomenon, various hydrophilic (water-based) clearing solutions were developed. However, water-based methods have had lower tissue clearing performance (Silvestri etਊl., 2016), while preserving the fluorescence. Figures 2 C and 2D illustrate preserving GFP fluorescence in embryonic heart of different stages (ED10.5 and ED14.5) by the CUBIC tissue clearing method. Besides using a hydrophilic clearing solution, another approach to maintain fluorescence is a tissue transformation method involving embedding the tissue in acrylamide gel𠅌LARITY (Chung etਊl., 2013).

Examples of immunostaining and preserving GFP fluorescence with tissue clearing on the developing mouse heart

(A and B) Immunohistochemistry combined with BABB on ED 9.5 mouse embryo, smooth muscle actin antibody (SMA) in red labeling the myocardium, CD31 (PECAM-1) in green staining the endocardium (B), and DAPI nuclear staining (not very distinct at this low magnification) in blue.

(C and D) Preservation of natural GFP fluorescence with CUBIC tissue clearing on ED 10.5 (C) and ED 14.5 (D) in mouse Connexin 40 - GFP (Cx40-GFP) embryo hearts with superimposed autofluorescence in red. Ventricular trabeculae and atria are positive for Cx40 at these stages. Autofluorescent blood is present in the ventricles (C). All images were captured with confocal microscopy. Scale bar represents 100 μm in all figures.

Detailed analysis of the fluorescence preservation by various clearing methods was performed by Xu and colleagues (Xu etਊl., 2019b ) on intestinal tissues. They confirmed that most aqueous clearing methods performed better in fluorescence preservation than organic solvent-based ones. On measuring signal to background ratio after tissue clearing, they found that the best method to preserve fluorescence was FRUIT, followed by ScaleS and SeeDB. However, CUBIC and PACT preserved similar levels of fluorescence as 3DISCO and uDISCO, which are the organic hydrophobic clearing agents. They did not include CLARITY and its variations in their testing. Even though their study included heart tissue, the part about fluorescence preservation was performed on intestine, which may react differently from the dense, highly vascularized heart tissue. Therefore, further analysis of 3DISCO, uDISCO, and FRUIT on cardiac tissue is needed. An overview is summarized in Figureਁ .

Only few studies have directly analyzed compatibility of tissue clearing and fluorescence preservation in the adult or embryonic heart tissue. In our previous study (Kolesova etਊl., 2016) we used CLARITY, SCALE, CUBIC, and DBE clearing methods and compared their GFP fluorescence preservation ability in embryonic hearts (illustrated in Figures 2 C, 2D and ​ and3B 3 B with CUBIC tissue clearing on Cx40:GFP trabeculae and coronary vasculature in embryonic hearts). We found that CUBIC cleared the tissue to a deeper level compared with SCALE and therefore was more suitable for analysis of intact hearts. However, although SCALE also preserved the GFP signal, it only cleared the superficial layers of the heart, which may be sufficient for studies of the great vessels of the coronary vasculature (Kolesova etਊl., 2016). The difference between SCALE and CUBIC was more obvious in postnatal hearts, where CUBIC effectively cleared all the way through the hearts, whereas SCALE did not (Shaikh Qureshi etਊl., 2016). Another study tested the ability of CUBIC to preserve various fluorescent signals in the heart tissue (Nehrhoff etਊl., 2016). They found that CUBIC clearing can also preserve other fluorescent proteins such as TdTomato and GFP in 750-μm-thick heart sections.

Coronary vasculature visualization in the developing hearts

(A) Vasculature was injected with DiI in a quail ED9 embryo.

(B) Coronary arteries were visualized on heart surface of an ED18.5 Cx40-GFP mouse embryo cleared with CUBIC. Imaged on a confocal microscope.

(C) Analysis of the DiI injected heart with coronary vessels pseudocolored to indicate depth within the ventricular wall, ED13 quail embryo.

(D) Juvenile mouse heart with coronary vasculature injected with Microfil and main coronary arteries indicated with color: Right coronary artery (orange) and its branch, septal artery (red), and left coronary artery (yellow). Imaged on a micro-CT scanner.

CLARITY also has been shown to preserve many fluorescent proteins such as GFP, mCherry, mOrange, and Cerulean in heart tissue (Sereti etਊl., 2018). Also, modified CLARITY protocols have been used to analyze cardiac tissue. SUT (Scheme Update on tissue Transparency, combination of CUBIC and CLARITY) has been used to analyze fibrotic healing in myocardial infarction (Wang etਊl., 2018). SCM (Simplified CLARITY Method) has been used to analyze heart tissue (Sung etਊl., 2016) however, better results were obtained when the blood was washed from the heart prior to fixation as decolorizing of heme using aminoalcohol treatment reduced YFP fluorescence in heart samples (Sung etਊl., 2016).


Cell lines

Human breast cancer cell lines MCF7 (HTB-22, ATCC) and MDA-MB-231 (HTB-26, ATCC) were maintained in α-minimum essential medium (α-MEM SH30265, GE Healthcare), supplemented with 10% fetal bovine serum (FBS S11150, Atlanta Biologicals) and 1% penicillin/streptomycin (P/S 15140, GIBCO). The PC3 human prostate cancer cell line was maintained in RPMI–1640 medium (23400–021, GIBCO) supplemented with 10% FBS and 1% P/S. Head and neck squamous cell carcinoma (HNSCC) OSC19 and HN5 cells were maintained in Dulbecco’s modified Eagle medium (DMEM SH30022FS, GE Healthcare), supplemented with 10% FBS, 1% P/S, 1 mM sodium pyruvate (11360070, GIBCO), 2 mM L-glutamine (25030081, GIBCO), 1% MEM vitamin (11120052, GIBCO), and 1% MEM non-essential amino acids (NEAA 11140050, GIBCO). The HCT116 human colorectal carcinoma cell line and its derived cells – DNA-PKcs knock-out (KO) cells 41 – were maintained in α-MEM supplemented with 10% FBS and 1% P/S. All cells were cultured in a humidified 5% CO2 incubator at 37 °C.

Fabrication of PDMS-HDA device

The fabrication of the PDMS-HDA device relied on a flat substrate (i.e. 22*22 mm 2 cover glass) fastened with an adhesive tape (3 M scotch 810), which is used as a mold to prevent leakage of the solution Fig. 1(a). Polydimethylsiloxane (PDMS Sylgard 184, Dow Corning) mixture was then poured into the mold and cast at 125 °C for 1 h. The weight ratio of the base to the curing agent was 10:1, and the weight of the PDMS prepolymer was 0.5 g per chip. The chip was sterilized by UV for 30 min after removing the tape and stored in a sterilized dish by sealing with parafilm at room temperature until use.

Three-dimensional spheroid culture by PDMS-HDA

Before the cells were loaded, a 1 ml type I collagen solution (

4.26 mg/ml C3867, SIGMA) was prepared by adding cell culture medium and 1 N sodium hydroxide for pH adjustment (details are reported in the Supplementary Table S1). Before each individual experiment, the pH of the collagen solution was measured to be around pH 7.4. The cells were then resuspended in solutions with different concentrations of collagen (0, 50, 500, and 1000 μg/ml). In 3D multicellular spheroid culture Fig. 1(c), cellular drops with a volume of 1 μl were dispensed onto the PDMS-HDA device (step 1) by a liquid handing machine (Versa 10 spotter, Aurora Instruments Ltd.) Fig. 1(b). Each drop had an average diameter of 1.4 mm. The device was flipped and placed in a 6-cm cell culture dish, which had been pre-filled with medium of 750 μl and sealed with parafilm to prevent evaporation of the cell-containing drops. The whole set was transferred immediately to a humidified incubator at 37 °C overnight to allow the formation of collagen fibrils, which contributed to the sedimentation and aggregation of the cells (Steps 2 and 3), promoting the generation of cellular spheroids (Step 4).

Evaluation of cell spheroid size

The mean volume of the 3D-cultured cell spheroids was calculated based on the measurements of their diameters by an open-source imaging software (Fiji). Although some spheroids were oval or shaped irregularly, the mean diameter (l) was determined by the following equation: l = (a × b) 1/2 , where a and b represent the two orthogonal diameters of each spheroid. The mean volume (V) of the spheroids was then evaluated by the equation (V=4 imes pi imes <(l/2)>^<3>/3) .

Cell labeling and imaging

Cultures on the PDMS-HDA device were stained with a mixture of 2 μM calcein AM and 4 μM ethidium homodimer-1 (live/dead viability kit, L3224, Thermo Fisher Scientific Inc.) to stain for live and dead cells, respectively. The device was incubated under 5% CO2 at 37 °C for one hour. Bright field and fluorescent images were captured using a CCD camera (SPOT Flex, SPOT Imaging) on an upright microscope. Image analysis was conducted by the Fiji imaging software.

Drug screening assay

To demonstrate that the PDMS-HAD device can screen for anti-cancer drugs, different tumor spheroids (of MCF7, MDA-MB-231, PC3, OSC19, and HN5 cells) were tested after treatment with the chemotherapeutic drugs paclitaxel (T7402, SIGMA) and cisplatin (P4394, SIGMA), at concentrations of 0 to 50 μg/ml. Based on the procedure described earlier, single cell spheroids were obtained at an initial cell density of 450/μl within each 500 μg/ml collagen-containing medium drop for a 2-day culture. Following the procedure shown in Fig. 2(a), a 1-μl drug-containing drop with two times the final testing concentration (0, 5, 25, or 50 μg/ml) was dispensed to the appropriate cell-containing drops, resulting in a total volume of 2 μl for each drop. After a 2-day drug treatment, cell viability was achieved using the CellTiter-Blue assay (at 520 nm excitation and 590 nm emission G8080, Promega Corp.) with a volume of 1 μl per drop cells were incubated for 4.5 h. A fluorescent microscope (BX51, OLYMPUS) was used to capture the fluorescent images of the cell-drug-containing drops. Fluorescent images were used to interpret the quantity of live cells. The fluorescent intensities of each drop were then analyzed by the Fiji imaging software. The relative cell viability, corresponding to the fluorescent intensity, was normalized against the untreated cells under different drug concentrations.

For on-device 2D drug screening, 1 μl of the cell suspensions mixed with 2 μg/ml collagen, used to facilitate the adhesion of the cells on PDMS, were first dispensed onto the device and cultured for 2 days without flipping. The following drug sensitivity experiments were conducted under the same conditions as the 3D drug screening mentioned earlier. For 2D control experiments from 96-well plates, cells were plated at a density of 6000 cells in 100 μl medium per well for 2 days. The media were exchanged accordingly with drug-containing media for an additional 2-day culture. Ten microliters of CellTiter-Blue were added to each well and the plates were incubated for 4.5 h. A plate reader (POLARstar Optima, BMG LABTECH) was used to determine the fluorescent intensity, and cellular viability was evaluated following the procedure described earlier.

Tumor dissemination assay

The tumor dissemination assay was conducted following the procedure shown in Fig. 3(a). Tumor spheroids cultured for 2 days were generated from an initial cell seeding of 900 cells. The spheroid-containing drops were dispensed with 10 μl medium drops with or without the drugs. The resulting height of each drop (a total volume of 11 μl) was around 1.2 mm. For the combined treatment, drugs were added to the device before γ-ray irradiation. The device was flipped and placed on a 6-cm cell culture dish. The supported wall of the device had a height of 1.5 mm and sufficiently prevented the unwanted contact between the drop array and the examined substrate. The spheroid array was shifted to the substrate within seconds, followed by exertion with uniform pressure on the device, simultaneously separating it from the substrate see the example in Fig. 3(b). The volume of each shifted drop was around 5 μl. The dish was filled with the medium and sealed with parafilm to prevent evaporation. The dish was transferred to a 5% CO2 incubator at 37 °C overnight to allow the cell spheroids to adhere to the substrate. Medium with or without the drugs was added to the dish after 1 day. Cellular dissemination was observed daily under the microscope, and the migration area was derived by measuring cellular migration subtracted from the initial area at day 0.

Three-dimensional co-culture assay

The PDMS-HDA device was adapted to the procedure shown in Fig. 4a to study whether two individual cell cultures in 3D could influence each other. The device is an emerging tool for modeling cell-cell interactions at different tissue levels 32 . Two cell spheroids (e.g. cell type A and type B) were generated at a gap of 1.5 mm. After culturing for 2 days, 1 μl of Matrigel was dispensed onto the cell-containing drops manually at 4 °C. Ten microliters of medium were added to the two adjacent drops after the gelation of Matrigel at 37 °C overnight. The mean diameters of two separate spheroids were observed daily. The spheroid volume was evaluated by the equation mentioned earlier, (V=4 imes pi imes <(l/2)>^<3>/3) .

Three-dimensional invasion assay

To perform the tumor spheroid invasion assay with the PDMS-HDA device, cell spheroids were first generated on the device for 24 h, following the procedure shown in Fig. 5(a). Three-dimensional scaffold matrices of 10 μl were added to each cell spheroid-containing drop, which was mixed with Matrigel and collagen-I (in PBS at pH 7.4) at a ratio of 1:1. The final concentration of Matrigel was 5 mg/ml and the concentration of collagen-I was 1.5 mg/ml. The addition of collagen-I to Matrigel facilitated the invasion by increasing matrix stiffness 42 . We observed that the cells in the mixed gel invaded more effectively than Matrigel or collagen gel used alone (data not shown). Five microliters of epidermal growth factor (EGF 236-EG-200, R&D systems)-encapsulated scaffold matrices were dispensed beside the corresponding spheroid drop at a gap of 1.5 mm. The EGF was used at concentrations of 0, 10, and 50 ng/ml. After gelation of the scaffold matrix at 37 °C for 1 h, 20 μl of standard culture media or media containing NU7441 were dispensed and connected with the two adjacent gel drops at day 0. The addition of the medium allows the generation of an in vitro-like microenvironment, in which a gradient of EGF from the encapsulated matrix could be generated to stimulate and coordinate nearby spheroid invasion Fig. 5(a). For the combination treatment, drugs were added to the device before γ-ray irradiation. Spheroid invasion was observed daily by microscopy. In addition, the invasion area was evaluated as a measurement of cellular spreading subtracted by the initial area at day 0.

Statistical analysis

Student’s t test was used to compare data from two groups. The one-way TukeyHSD ANOVA test was used to compare data from more than two groups. A p < 0.05 was considered to be statistically significant.

Author contributions

TN and MS designed research TN, ME, CR, JS, JMU, PF, SW, LO and AM performed research PL, MDF, BMM, AC and AJM contributed new reagents/plant lines/analytic tools TN, ME, CR, JS, PF, SW and SJM-S analysed data and TN and MS wrote the paper with assistance from all co-authors.

Please note: Wiley Blackwell are not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

Fig. S1 Purification of heterologous roGFP2-Orp1 protein, emission spectra and H2O2-dependent oxidation rates.

Fig. S2 Analysis of cytosolic roGFP2-Orp1 redox state in different Arabidopsis seedling tissues.

Fig. S3 Redox responses of independent roGFP2-Orp1 Arabidopsis lines and in planta spectra of reduced mitochondrial sensor.

Fig. S4 Expression of the fluorescent probe cyt-roGFP2-Orp1 in Arabidopsis.

Fig. S5 Redox response of 2′,7′-dichlorodihydrofluorescin diacetate and mt-roGFP2-Orp1 in Arabidopsis seedlings to menadione.

Fig. S6 The response of cyt-roGFP2-Orp1 to flg22-triggered oxidative burst in diphenylene iodonium-treated Arabidopsis leaf discs.

Methods S1 Cloning of sensor constructs and generation of plant lines.

Methods S2 Plant culture.

Methods S3 Purification of roGFP2-Orp1 protein.

Methods S4 Elicitors.

Table S1 Primer sequences for cloning of the expression vectors.

Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.

Fluorescent Mouse Reporter Models for Tissue Imaging

Several types of fluorescent reporter models, depending on the application both constitutive and inducible, have been used individually or in combination for live ectodermal tissue imaging (summarized in Table 1). These models can be divided into three, partially overlapping, categories: indicators of cellular behaviors, cell identity/fate, signaling activity. Examples of reporters for cellular behaviors include reporters to visualize cell divisions such as the R26-H2B-EGFP model (Hadjantonakis and Papaioannou, 2004) and the ubiquitinoylation oscillator based fluorescent (Fucci) cell cycle reporters that allow direct real-time follow-up of the progress of the cell cycle in individual cells with a nuclear, dynamically color changing, fluorescent signal (Sakaue-Sawano et al., 2008 Mort et al., 2014). These reporters have also been used to follow cell movements with live imaging. Fluorescent inducible cell membrane bound reporters such as the R26R mT/mG (Muzumdar et al., 2007) and R26R Confetti (Snippert et al., 2010) allow visualization of tissue type, cell shape changes, lineage tracing, and scarce labeling approaches tracking of individual cells. The signaling reporter TCF/Lef:H2B-GFP (Ferrer-Vaquer et al., 2010) is a sensitive fluorescent reporter of canonical Wnt signaling activation and is based on tandem transcription factor binding sites driving expression of the H2B-EGFP fusion protein. Examples of fluorescent reporters for cell identity are the Shh GFP𠄼re (Harfe et al., 2004) reporter visualizing signaling center identity, Sox2-GFP for hair follicle dermal condensate cells (Biggs et al., 2018) and keratin 17-GFP to visualize the ectodermal organ epithelium (Bianchi et al., 2005 Ahtiainen et al., 2014).

Table 1. Fluorescent mouse reporter models for live tissue imaging of ectodermal embryogenesis.


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Problem 1

Sections detaching from charged glass slide (step 3).

Potential solutions

When performing vertical washes (in glass Coplin staining jars), it is possible that tissue sections start to detach. To prevent this, numerous approaches can be taken into consideration:

Solution 1: While cutting the sections in the cryostat and placing them onto the glass slides, they should be kept at room temperature for 30 min to 1 h to promote complete adherence of the tissue section to the glass.

Solution 2: Slides such as APES (aminopropyltriethoxysilane) treated glass slides may be used (instead of SuperFrost™ Plus slides) to increase tissue adherence (

Solution 3: Before starting the staining, enough time should be given for the slides to be at room temperature, to ensure that the sections are completely attached to the glass slides.

Solution 4: If tissue is not already fixed, fix with PFA on slide to promote further adhesion.

Solution 5: While performing the staining, gently replace the buffer between washes and, when using shaking, select a low speed.

Solution 6: If the sections continue to detach, carry out all washes horizontally in a drop-wise fashion.

Problem 2

High fluorescence signal is detected in sections that were incubated with the haptenic sugar (inhibitory controls) (step 14 or 25).

Potential solution

If lectin binding is carbohydrate-mediated, by incubating a lectin with its haptenic sugar, the haptenic sugar will occupy the lectin-binding site and prevent the lectin from binding to the glycans in the tissue, leading to significantly decreased signal in those sections (complete abolition should not always be expected). Consult manufacturer’s instructions to confirm the most appropriate sugar to use. Additionally, it is worth noting that some lectins can only be inhibited by complex sugars and not monosaccharides, in which cases appropriately glycosylated glycoproteins should be employed (Brooks and Hall, 2012). If lectin binding is not significantly decreased in the presence of the haptenic sugar, lectin binding is either non-specific or mediated by the non-carbohydrate binding sites present in the lectin (Gerlach, etਊl., 2012). A different lectin with similar desired specificity should be selected in the latter case.

Problem 3

High background staining/fluorescence signal (step 14 or 25).

Potential solution

If a high background staining is detected, there are several steps that can be tried to reduce it. This can be due to several reasons:

Reason 1: Insufficient blocking of non-specific binding sites, which can be overcome by increasing the concentration of periodate-treated BSA or by extending the incubation period of the blocking step.

Reason 2: Excessive lectin concentration, incubation temperature, or incubation time. In these cases, it is advised to reduce the concentration of lectin and/or to incubate sections at 4ଌ instead of at room temperature and/or for a reduced period of time. Also, the lectin used could be potentially diluted in a buffer containing the blocking agent (e.g., 0.1% periodate-treated BSA in TBS-T or TBS). The use of other blocking agents (such as normal serum, which is frequently used in immunohistochemistry) is not appropriate as it possesses multiple glycosylated molecules, which might inhibit specific binding of lectins to the tissue or cause non-specific binding.

Reason 3: Number and duration of washes are too short. These can be increased to ensure that any lectin that is non-specifically bound is removed.

Reason 4: The tissue might have physiological structures/molecules that can cause auto fluorescence due to endogenous fluorophores (e.g., lipofuscin) (Croce and Bottiroli, 2014). To solve this problem, it is crucial to know the features and composition of the tissue and try to address them by finding a specific compound that can bind to these structures and reduce their auto fluorescence.

Problem 4

No fluorescence signal is detected in the stained tissue sections (step 14 or 25).

Potential solution

Lack of fluorescence signal in the samples might be caused by multiple factors:

Factor 1: Not enough lectin is bound to glycans of interest, for which the recommended course of action would be to use a higher concentration of lectin, to incubate it during a longer period (e.g., 2 h or even 3 h) or to incubate it at a higher temperature (e.g., at 37ଌ).

Factor 2: The lectin might have been improperly stored (e.g., kept at room temperature for long periods or in an environment not protected from light), which could lead to quenching of its signal or loss of its function. To ensure that the lectin has kept its function and labeling, a positive control should be run in parallel in every staining experiment. This positive control should be a tissue known to express the glycosylation motif specific to that lectin as characterized by the literature or based on previous in-house experiments.

Factor 3: The specific glycosylation motif might not be present in the tissue sample. To confirm this, a positive control using a sample where this glycosylation motif has been reported to be found should always be run in parallel.

Factor 4: The fixation procedure may modify the glycan structure that is necessary for recognition by the lectin. To solve this issue, a shorter fixation period or a different fixation method are recommended.

Factor 5: TBS buffer might be contaminated with bacteria, which might alter its properties and compromise the staining and functions of the lectin. For this, it is advised to only use freshly prepared sterile-filtered or autoclaved TBS.

Problem 5

The positive signal associated with the glycan distribution is not homogenous across the tissue section (step 14 or 25).

Z.Y. and J.L. contributed equally to this work. Conceptualization — X.C., X.Z., and H.W.O. Supervision — X.C. and H.W.O. Investigation — Z.Y., J.L., W.Z., M.L., R.L., B.Z., Y.C., and Y.H. Formal Analysis — J.L. and R.Y. Methodology — K.Z., C.F., and C.A. Writing–Original Draft — Z.Y. and J.L. Writing–Review & Editing — A.E., J.Z., and H.W.O. Funding Acquisition — X.C., B.W., and H.W. All authors read and approved the manuscript.

Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.