Histochemical Techniques in Plant Science: More Than Meets the Eye

Abstract

Histochemistry is an essential analytical tool interfacing extensively with plant science. The literature is indeed constellated with examples showing its use to decipher specific physiological and developmental processes, as well as to study plant cell structures. Plant cell structures are translucent unless they are stained. Histochemistry allows the identification and localization, at the cellular level, of biomolecules and organelles in different types of cells and tissues, based on the use of specific staining reactions and imaging. Histochemical techniques are also widely used for the in vivo localization of promoters in specific tissues, as well as to identify specific cell wall components such as lignin and polysaccharides. Histochemistry also enables the study of plant reactions to environmental constraints, e.g. the production of reactive oxygen species (ROS) can be traced by applying histochemical staining techniques. The possibility of detecting ROS and localizing them at the cellular level is vital in establishing the mechanisms involved in the sensitivity and tolerance to different stress conditions in plants. This review comprehensively highlights the additional value of histochemistry as a complementary technique to high-throughput approaches for the study of the plant response to environmental constraints. Moreover, here we have provided an extensive survey of the available plant histochemical staining methods used for the localization of metals, minerals, secondary metabolites, cell wall components, and the detection of ROS production in plant cells. The use of recent technological advances like CRISPR/Cas9-based genome-editing for histological application is also addressed. This review also surveys the available literature data on histochemical techniques used to study the response of plants to abiotic stresses and to identify the effects at the tissue and cell levels.

Introduction

Histochemical staining has become a widely used technique in the field of plant science for the localization of proteins, enzymatic and promoter activities, elements, oxidative stress markers, minerals and cell wall components in plant cells/tissues (Al Bitar et al. 2002, Romero-Puertas et al. 2004, Law and Exley 2011, Prabhu et al. 2011, Hassan and El-Awadi 2013, Giuliani et al. 2018, de Ávila et al. 2019). Histochemical methods are also beneficial for the in situ quantification of several metabolites, allowing to determine the place of their respective production or action (Dubey and Trivedi 2012). Histochemistry allows in situ analysis of specific components while retaining the 3D spatial information of the cell/tissue by using particular staining and photographic recording under microscopes, such as light, electron, stereoscopic, fluorescent and confocal microscope (Hassan and El-Awadi 2013). The histochemical detection is based on the simple principle that when a tissue or cell is exposed to a staining solution, it will react specifically by forming a colored insoluble end product detectable by microscopy (Ruifrok and Johnston 2001). Different microscopes can be used depending on the specific characteristic of the compound. For instance, light microscopy is used to image the birefringence of compounds like starch grains or crystals (Carlsson 2007). Fluorescence microscopy is used to image compounds that emit fluorescence when irradiated with a certain wavelength. In this case, the compound to be analyzed can be fluorescent or react with a specific fluorescent probe (Lichtman and Conchello 2005, Masters 2010). Confocal microscopy allows the study of compounds and organelles in vivo in the X, Y, Z axes and even analyze the dynamics of metabolites and organelles in the tissue over time (Rodríguez-Serrano et al. 2009). Electron microscopy allows the visualization and measurement of biomolecules at the subcellular level, although in this case, tissue fixation is needed, and in vivo studies are not possible (Bozzola and Russell 1999). Although used for decades, histochemistry remains a reliable complementary tool enabling physiological studies in different experimental conditions. In the field of plant science, histochemical analyses nicely complement analytical approaches, such as the biochemical quantification of (macro) molecules and enzymatic activities, by providing data concerning the localization of promoter activity, subcellular mapping of proteins, metabolites, elements and minerals. Such data are particularly relevant in stress physiology since they enable linking detection with spatial information at the cellular/tissue level. We propose to use histochemistry, a technique that is not too expensive nor too demanding in terms of analytical procedure, as an initial analytical tool to choose a specific experimental set-up or time-points that will then be investigated in more detail using high-throughput studies, such as -omics and (bio)chemical analyses.

Localization of Metals

The cellular localization of metals can be achieved with the help of various metallochrome indicators and several low specific fluorescent dyes to detect several metals, namely cadmium (Cd), copper (Cu), iron (Fe), mercury (Hg), nickel (Ni), lead (Pb), strontium (Sr) and zinc (Zn) (Seregin et al. 2011, Seregin and Kozhevnikova 2011). Histochemistry can be especially useful to study metal transport across cellular membranes of plants and to unveil the strategies used by plants to cope with metal toxicity. The uptake, translocation and accumulation of heavy metals also differ according to the metal and tissue type (Clemens 2006; Table 1). Several specific and nonspecific staining methods have been reported for different metals and are briefly discussed in the next sections.

Localization of Zn

Zn is a crucial micronutrient for plants and a cofactor of many enzymes related to photosynthesis, nitrogen metabolism, synthesis of auxin, nucleic acids and proteins (Rout and Das 2009, Seregin et al. 2011). From the soil solution, it primarily enters the plant through roots as Zn²⁺, which forms a complex with organic ligands (Martin et al. 2007, Kenderešová et al. 2012). Excess Zn becomes toxic as it impairs growth by affecting morphogenesis (Sresty and Rao 1999, Rout and Das 2009). In the opposite scenario, Zn deficiency also negatively affects plant growth by causing root apex necrosis, chlorosis, bronzing and imbalances in auxin physiology, ultimately resulting in internode shortening, epinasty and leaf size reduction (Martin et al. 2007). Its accumulation in plants can be detected by using the metallochromic indicator Zincon and the lipophilic fluorescent indicator Zinpyr-1, imaged under light, fluorescence or confocal microscopes. Zincon produces a blue-colored complex with Zn and has high selectivity toward this metal (Seregin et al. 2015; Table 1, Fig. 1A), while Zinpyr-1 produces a green fluorescence signal after reacting with Zn (Seregin et al. 2015; Fig. 1B). It can also be detected by using different fluorescent indicators, such as Zinquin, RhodZin-3, FluoZin-1, FluoZin-2, FluoZin-3, IndoZin-1, FuraZin-1, TSQ, Newport Green PDX and Newport Green DCF (Haugland 2002). These dyes differ in their affinity and specificity for Zn and their plasma membrane permeability (Seregin and Kozhevnikova 2011). A previous study has shown that Zn excess in maize (Zea mays L.) seedlings affects root growth, cell division and cell elongation (da Cunha and do Nascimento 2009). Furthermore, Seregin et al. (2011) analyzed the high concentration of Zn located in the root tips and the apoplast of most plant organs. Zn was also detected by staining Arabidopsis roots with Zinpyr-1. Sinclair et al. (2007) used a specific Zn chelator N,N,N′,N′-Tetrakis (2-pyridylmethyl) ethylenediamine as a negative control, which allows somewhat to differentiate between autofluorescence and specific Zn-derived fluorescence. Positive controls prepared by incubating the plants with high Zn concentrations are useful to confirm the metal location (Sinclair et al. 2007). A similar approach was followed by Seregin and Kozhevnikova (2011) to analyze Zn distribution in Thlaspi arvense roots.

Localization of Cd

Cd is one of the highly toxic heavy metals, the source of Cd in the environment is different anthropogenic activities that pollute soils and it further bio-accumulates in the food chain (Dorne et al. 2011, Vaculík et al. 2012). In higher plants, the level of Cd uptake is determined by the Cd concentration in the soil and the physical and chemical properties of the soil. Several studies carried out in different plant species demonstrated adverse effects of Cd on root and shoot growth, water and nutrient uptake, photosynthesis, hormone balance and reactive oxygen species (ROS)-derived oxidative damage (Sandalio et al. 2012). However, the effect can differ in different plant species, depending on the amount of metal and period of treatment (Sandalio et al. 2012). Cd detection has been carried out using dithizone (Fig. 1G), although this stain is not just specific to Cd and can also react with other metals, such as Pb (Seregin and Ivanov 1997, Peco et al. 2020b). However, it has mostly been used to study Cd accumulation in different parts of plants, particularly in the root, as it is an organ that accumulates high levels of Cd (Xu et al. 2018, Peco et al. 2020a). According to a study on maize seedlings, Cd was indeed found only in vascular tissues and adjacent parenchyma cells in the roots (Wójcik and Tukiendorf 2005). In Populus canescens, Cd was detected in the phloem of the bark (primarily in vacuoles of phloem cells), on the root surface and in the apoplastic region of the root cortical cells (He et al. 2013). Leaves stained with Cd-dithizone showed Cd localization in different regions, notably in leaf veins, mesophyll cells around veins, leaf trichomes and petioles region. These results were corroborated by an energy-dispersive X-ray. All these data suggest a preferential accumulation of Cd along with the plant vascular system (He et al. 2013), although Cd can also be accumulated in trichomes (Peco et al. 2020a). Dithizone staining has also been helpful to study the effects of different compounds, such as melatonin and silicon, on plant response to this metal.

Localization of arsenic

Arsenic (As) is a toxic metalloid, naturally existing in the environment due to volcanic explosion and metal ores. Anthropogenic sources of As pollution are fossil fuel combustion, timber preservatives and use of pesticides. Arsenic exists in the environment in different forms, such as arsenite (AsIII), arsenate (AsV), monomethylarsonic acid, dimethylarsinic acid, arseno-betain and arseno-choline (Tangahu et al. 2011, Sharma 2012, Souri et al. 2017). As accumulation induces oxidative stress in plants that reduces productivity by interfering with various metabolic pathways (Souri et al. 2017). It also induces oxidative stress during the conversion of AsV to AsIII (Sharma 2012) and through the disruption of electron transport chains in mitochondria and chloroplasts, activation of glycolate oxidase associated to photorespiration, antioxidant inactivation and glutathione (GSH) depletion (Souri et al. 2017). Moreover, As toxicity leads to root hair loss, damage of epidermal cells, cortex and thylakoid membrane, ultimately resulting in cell death (Sharma 2012). Furthermore, As causes disorders in plant physiology by impairing gas exchange in leaves, lack of nutrients availability, disruption in water potential, decrease in protein content, inhibition of phytase activity and chlorophyll biosynthesis and also decrease in photosynthetic efficiency and biomass (Sharma 2012). As is mainly accumulated in shoots in hyper-accumulator plants, such as the fern genus Pteris (Sridhar et al. 2011, Souri et al. 2017). The staining with CuSO₄ in Pteris vittata shows that As precipitates surrounding the stem vascular bundles (Sridhar et al. 2011; Fig. 1C). However, to study the speciation and distribution of As, other approaches have been used, such as synchrotron-based X-ray absorption, X-ray absorption near-edge structure spectroscopy (XANES), spatially resolved microspectroscopic techniques, μ-X-ray fluorescence (μ-XRF) and particle-induced X-ray emission (Lombi et al. 2009).

Localization of chromium

Chromium (Cr) is known as toxic heavy metal for plants. Cr exists in nature in two forms, such as trivalent Cr (III) and more toxic hexavalent Cr (VI) (Shanker and Pathmanabhan 2004). In plant tissue, Cr (VI) reduced to Cr (III). Cr being a redox metal, it induces oxidative stress in plants. It can also affect the antioxidant activities of superoxide dismutase, catalase, and glutathione reductase (Shanker and Pathmanabhan 2004). Chrome Azurol S staining is used for the localization of Cr giving rise to blue precipitates in plant tissues (Sridhar et al. 2011; Fig. 1E). In this way, Sridhar et al. (2011) have observed that the brake fern (P. vittata) accumulates high Cr concentration in roots, while Shanker and Pathmanabhan (2004) found that in plants Cr uptake and translocation from root to shoot is narrowed because of the tendency of Cr (III) to bind to the cell wall. Some of the symptoms of Cr toxicity are the decrease in intercellular spaces, shrinkage of epidermal cells, parenchyma and water stress disturbances (Shanker and Pathmanabhan 2004, Sridhar et al. 2011) (Table 1). μ-XRF imaging and XANES analysis have also been used to analyze Cr distribution in Typha angustifolia (lesser bulrush), where the metal was observed mainly associated with vascular tissues in the root (Vidayanti et al. 2017).

Localization of Ni

Ni is known as an essential micronutrient for an optimal plant growth. However, it is strongly phytotoxic at a higher concentration by promoting the oxidative stress as a result of ROS production and disturbances of antioxidant defenses. Generally, Ni causes reduction in plant growth and also accelerates chlorosis, necrosis and wilting (Aydaş et al. 2013, Bhalerao et al. 2015). Dimethylglyoxime (DMG) staining is particularly well suited for Ni localization. The complexes Ni-DMG can be imaged in the light and fluorescence or confocal microscope (Gramlich et al. 2011). The images are recorded in the three color channels, red, green and blue, to discriminate DMG from non-DMG signals. In this way, it is possible to discriminate DMG-dependent signals from cell autofluorescence (Gramlich et al. 2011). Sodium rhodizonate can also be used for the detection of Ni and Sr (Seregin and Kozhevnikova 2004, Seregin and Kozhevnikova 2011) (Table 1). Gramlich et al. (2011) studied Ni distribution in roots, stems and leaves of Berkheya coddii using DMG. However, the obtained results showed significantly higher Ni accumulation in leaves than in stems and roots. Therefore, it was concluded that Ni was not distributed homogeneously but accumulated in some parts of the leaves, for instance particularly in the epidermis (Table 1). Alyssum corsicum also shows higher Ni accumulation in shoots than in roots (Aydaş et al. 2013). Furthermore, this plant, when treated with Ni, shows the highest metal accumulation in the stomata of the adaxial surface and in the trichome rays of the abaxial surface.

Localization of Fe

Fe is a known vital microelement for plant growth. It partakes in multiple processes in plants, such as respiration, photosynthesis, oxygen transport and DNA synthesis (Kao 2014). Therefore, its nutritional status is vital for the cell metabolism and functioning and was reported to be necessary for keeping the balance between cation/anion ratio, synthesis of biomolecules and energy conservation in plant membranes (Rout and Sahoo 2015). Fe being a redox metal plays a prominent role in the proper functionality of redox-active proteins involved in respiration, photosynthetic mechanism and nitrogen fixation (Takahashi et al. 2009). It also participates in the citric acid cycle, sulfur and nitrogen assimilation and chlorophyll biosynthesis (Takahashi et al. 2009). The stains ‘Prussian blue’ and ‘Toluidine Blue O’ (Basic blue; Methylene Blue T50 or T Extra) have been used for localizing Fe in endospermic aleurone cells and scutellar cells of the rice plants (Krishnan et al. 2001; Table 1). Probably, the most widespread method used to detect Fe in plant tissues is the Perls’ staining, and DAB (3,3′-Diaminobenzidine) intensification technique (Fig. 1F), which has been successfully used to study Fe distribution in Arabidopsis and pea plants (Roschzttardtz et al. 2009, Roschzttardtz et al. 2013). Roschzttardtz et al. (2013) have performed an analysis of Fe distribution in different organs of Arabidopsis thaliana wild-type and mutant genotypes. Arabidopsis plants, both wild type and the vacuolar Fe transporter knockout mutant vit-1, accumulated Fe in the apoplast of the central cylinder in roots, while in iron-treated plants, Fe was accumulated in vascular tissues and in the chloroplast of surrounding mesophyll cells, mainly as a Fe-ferritin complex. Pollen grains accumulated Fe in amyloplasts (Roschzttardtz et al. 2013), as the vacuole is the iron storage compartment in the Arabidopsis embryos. By using Perls’/DAB and 4′,6′-diamido-2-phenylindole (DAPI) staining of the nucleus, Roschzttardtz et al. (2011) demonstrated the accumulation of Fe in the nucleolus of pea plants’ embryos. The unexpected presence of Fe within the nucleus and in the subcompartment nucleolus was identified through elemental imaging via microparticle-induced X-ray emission and X-ray fluorescence (Table 1).

Localization of Cu

Cu is an essential element and, being a redox-active transition metal, it is found to be involved in numerous physiological functions and mechanism in plants, such as photosynthesis, respiration, nitrogen fixation and pathogen defense, and in excess, it can also inhibit the plant growth (Shingles et al. 2004). Phen Green™ SK and Phen Green FL dyes have been used to detect Cu in plant cells (Seregin and Kozhevnikova 2011). Changes in the level of labile Cu in canola roots were detected by the Phen Green™ SK, an acetate ester that forms through the reduction by cell esterases and quenched by Cu ion binding (Zlobin et al. 2017). In the same way, the Phen Green™ SK (Dipotassium Salt) was used in the detection of the Cu absorption pathway in Saccharomyces cerevisiae (Sun et al. 2016). Shingles et al. (2004) analyzed the movement of Cu²⁺ in the thylakoid membrane of pea by loading the membrane with the Cu⁺-sensitive fluorophore Phen Green™ SK, which possess metal-binding phenanthroline covalently attached to fluorescein. The Cu distribution was located at the surface of the yeast cell by using PhenGreen™ (dipotassium salt and diacetate) and observed under the atomic force microscopy at 505 nm (Zu et al. 2006).

Localization of Pb

Pb is a highly invasive and lethal heavy metal reported with detrimental effects on seed germination, root growth, seedling development and transpiration (Pourrut et al. 2011). According to several studies, it can also induce ROS generation that subsequently causes oxidative stress in plants, thereby damaging plant cells and physiology (Pourrut et al. 2011). In the plant tissue, Pb can be histochemically detected by using sodium rhodizonate, which forms a bright scarlet-red precipitate with Pb but only at pH 3. The affinity of sodium rhodizonate to Pb is much lower than Sr (Seregin and Kozhevnikova 2011; Table 1). However, Pb can also be detected using dithizone as proposed by Comitre and Reis (2005). According to these authors, dithizone reacts with Pb to form complexes, while the chromogenic reagent dissolves in the organic phase and, therefore, the compound becomes dependent on Pb ion migration from the aqueous to the organic phase. Dithizone has been recently used to determine the localization of Pb in the root vacuoles and cell walls of the Pb-tolerant plant Biscutella auriculata (Peco et al. 2020b).

Localization of Plant Macronutrients

Sub-optimal availability of nitrogen (N), phosphorus (P) and potassium (K) is widespread over the world. N is an essential plant macronutrient mostly involved in the biosynthesis of nucleic acid, amino acids and proteins. Its deficiency causes growth reduction and decreased crop productivity (Rubio-Wilhelmi et al. 2011). Plants absorb nitrogen in the form of inorganic nitrate (⁠${\mathrm{N}\mathrm{O}}_3^{-}$⁠) that can be reduced to nitrite (⁠${\mathrm{N}\mathrm{O}}_2^{-}$⁠). Rhodamine 110 has been used as an important fluorescence quencher for the detection of trace nitrite in plants (Spence and Johnson 2010). Other elements, such as Ca and K, can also be detected by histochemistry. For instance, Krishnan et al. (2001) analyzed Ca²⁺ and K⁺ accumulation in rice plants by using Alizarin Red (used in bright-field and fluorescence microscopy) for the study of Ca²⁺-associated phytin granules and cobaltinitrite reagent for the localization of K⁺ (Krishnan et al. 2001). Alizarin Red, however, shows low sensitivity and other non-ratiometric and more sensitive dyes to image Ca²⁺ have been developed, including Fluo, Rhod and Calcium Green-1. Between the ratiometric dyes used to image Ca²⁺, Fura-2 and Indo-1 are the most used dyes in plants (reviewed in Kanchiswamy et al. 2014). The efficient intracellular fluorescent K⁺ probes, in addition to cobalnitrite, is potassium-binding benzofuran isophthalate (PBFI), which has been successfully used to analyze K⁺ distribution in pollen, anthers and papillae during pollination and fertilization in barley (Rehman and Yun 2006).

P is another major essential micronutrient in soil (1 ppm). Plants uptake P in the form of ${\mathrm{H}\mathrm{P}\mathrm{O}}_4^{2-}$ or H₂${\mathrm{P}\mathrm{O}}_4^{-}$ (soluble ionic phosphate) (Rodríguez and Fraga 1999). P is found abundantly in the topsoil strata. Low P availability induces changes in roots of annual crop plant that boost topsoil foraging (Lynch and Brown 2001). Excess Al and Mn availability, P deficiency and less availability of base, such as Ca²⁺, make the soil acidic (pH below 5.5), while decrease in the availability of P and micronutrient transition metals, such as Fe, Cu, Mn and Zn, makes the soil alkaline (pH above 7.5) (Nguyen et al. 2010). P accumulates in several microorganisms in the form of inorganic polyphosphate (polyP), which is a polymer of phosphate associated with phosphoanhydride bonds (Kornberg et al. 1999). The levels of polyP were estimated by the histochemical staining by using Toluidine Blue O and polyphosphate kinase, which converts polyP to ATP, followed by the ATP-luciferase assay (Ezawa et al. 2004). Several indicators detect Cl⁻ via diffusion-limited collisional quenching such as 6-methoxy-N-(3-sulfopropyl) quinolinium, N-(ethoxycarbonylmethyl)-6-methoxyquinolinium bromide, 6-methoxy-N-ethylquinolinium iodide and lucigenin (Asakawa et al. 2004).

Localization of Minerals in Plant Tissues: The Example of Silica

Plant tissue can accumulate biominerals, which provide mechanical protection to the tissues and act as a deterrent against herbivores and phytophagous insects. An emblematic example is biogenic silica (SiO₂), which is present in plants in amorphous form and typically found in structures, such as trichomes, stomata and silica cells in monocots (Kumar et al. 2017, Guerriero et al. 2018, Guerriero et al. 2019, Guerriero et al. 2020). The deposition of silica provides an additional line of defense to the entry of pathogens (Wang et al. 2017), and hence studying its localization in plants is valuable to identify the sites of deposition during biotic stress. For example, in Cucumis sativus, silicon is found in papillae after Sphaerotheca fuliginea infection (Samuels et al. 1994), a finding strengthening the hypothesis of a mechanical role in plants.

The silicaphilic fluor PDMPO 2-(4-pyridyl)-5-((4–(2-dimethyl amino ethyl amino carbamoyl) methoxy) phenyl) oxazole is known for its affinity to silica and used to localize siliceous structures in different plants (Law and Exley 2011). This oxazole dye initially used to study pH shifts within organelles (Diwu et al. 1999) partially neutralizes the negative surface charge of silica between a specific pH range (4–9.5): the pyridinium form is stabilized by reacting with the negatively charged silica and a proton is shared with a siloxide group (Parambath et al. 2016). In horsetail, thale cress, rice and hemp, PDMPO was used to visualize silicified structures after acid digestion (Law and Exley 2011, Guerriero et al. 2018, Guerriero et al. 2019, Guerriero et al. 2020). It should be noted that PDMPO was also used in vitro to monitor callose-assisted silica precipitation from an undersaturated silicic acid solution (Law and Exley 2011). Recently, PDMPO was also used on fresh leaves of okra to study the subcellular distribution of silica (Pierantoni et al. 2017).

Mapping the Expression of Genes

Histochemical localization of promoter activity can be done by the widespread β-glucuronidase (GUS) assay. In the analysis, by using Triton X-100, the 5-bromo-4-chloro-3-indolyl β-D-glucuronide (X-gluc) promoter is located by a blue color in transformed plants expressing the gus-A gene (Zainudin et al. 2015; Table 2). Hassan and El-Awadi (2013) reviewed the use of the GUS assay in transient expression experiments. GUS staining performed on tissues of transgenic rice plant identified the promoter activity of OsVDAC2, encoding a mitochondrial outer membrane ion channel, revealed staining of actively growing lateral roots and in pollen grains. Furthermore, intense GUS expression indicated that the OsVDAC2 promoter was expressed in all plant organs, with however strong staining in developing pollen grains (Al Bitar et al. 2002; Table 2). To overcome the staining requirement, low resolution and time taking procedure in GUS, simpler alternative exploring naturally occurring or genetically modified fluorescent proteins have been developed. The green fluorescent protein (GFP) is one of such most widely used fluorescent protein tag, which shows bright green fluorescence (509 nm) when visualized under blue light (395 nm) (Ortega-Villasante et al. 2016). The protein has 238 amino acids and makes a β-barrel 11-stranded helix, where the fluorophore is located and undergoes a spontaneous cyclization and oxidation of the Ser65-Tyr66-Gly67 motif (Orm et al. 1996, Ortega-Villasante et al. 2016). The gene encoding GFP was initially cloned from jellyfish and later modified in several studies to create more efficient and small size variants. For instance, a library of Arabidopsis cDNAs was created and attached to the 3′ end of GFP. Afterward, the library was transformed into Arabidopsisen masse and the transformed library was further used for transgenic plants showing various subcellular localization patterns (Tanz et al. 2013). Several other similar proteins emitting different bright colors have been cloned from different organisms and also improved with genetic engineering tools. Such fluorescent protein tags provide numerous opportunities to visualize, even at low concentration, a protein of interest, as well as protein–protein interaction (via Bimolecular Fluorescence Complementation, BiFC). A tool kit has been developed to efficiently express multigenes tagged with several different fluorescent proteins in plants. The availability of an exhaustive catalog of fluorescent proteins with recent technological advances in microscopy provides an opportunity for efficient histochemical studies (Hanson and Köhler 2001, Ehrhardt 2003).

Advances in Hybridization Histochemistry

Hybridization-based approaches like fluorescence in situ hybridization (FISH) allows the direct mapping of DNA sequences (Pinkel et al., 1988, Rodríguez-Serrano et al. 2009). This technique uses a florescent probe that hybridize (binds) to complementary sequence of DNA enabling to study the specific DNA fragments or genes present on chromosome (Pinkel et al. 1988). In addition, FISH technique can be used to detect RNA molecules which helps in understanding spatial-temporal gene expression (Rodríguez-Serrano et al. 2009, Solõs et al. 2012). Under Cd exposure, the regulation of the pathogen-related protein PrP4A was observed in pea leaves by FISH (Rodríguez-Serrano et al. 2009). Solís et al. (2012) also used FISH and reverse transcription PCR for the analysis of DNA methyltransferase-like genes MET1 during pollen embryogenesis. The level of this gene was upregulated during pollen maturation, while after pollen reprogramming, it was downregulated (Solõs et al. 2012). FISH also allows the profiling of the housekeeping gene PP2A in Arabidopsis roots (Duncan et al. 2016). Khrustaleva and Kik (2001) used the Tyr-FISH to detect the target T-DNA sequences on plant (Allium cepa) metaphase chromosome. Another technological improvement CASFISH was first successfully demonstrated by Deng et al. (2015). This approach harnessing recent advances in the Cas [clustered regularly interspaced short palindromic repeats (CRISPR)-associated] protein-based genome-editing technique has made FISH more sophisticated and precise (Deng et al. 2015). The Cas9 protein can be targeted to any specific sequence by using a guide RNA (gRNA) which is a short sequence (20 bases) complementary to the target site (Deng et al. 2015). In CASFISH, Cas9 protein needs to be first tagged with HaloTag ligand, which can be covalently linked with any organic fluorescent dye (Deng et al. 2015). Such assembly, efficient to in situ target-specific sequence, provides an enormous opportunity to study DNA localization (Fig. 2). Similarly, another approach termed as CRISPRainbow has been developed by Ma et al. (2016), where gRNA is engineered in a way that binds to fluorescent proteins. Thereafter, as like CASFISH, gRNA drives the dCas9 at the target site, which efficiently labels the desired DNA fragment (Ma et al. 2016). Such Cas9-based improved histochemistry techniques will facilitate several domains of life sciences, including plant science.

Hybridization Histochemistry for RNA Localization

Targeting RNA using programmed Cas9 is possible similar to the DNA targeting using the dCas9 with approaches like CASFISH and CRISPRainbow (Nelles et al. 2016). The RNA targeting can be achieved by using an oligonucleotide having Protospacer Adjacent Motif (PAM) sequence termed as PAMmer (Nelles et al. 2016). The PAMmer hybridizes to the complementary RNA target which facilitate the binding of the gRNA and Cas9 complex. However, the gRNA can bind to DNA encoding the RNA molecule instead of the transcribed RNA, and to avoid such DNA targeting, a mismatch can be introduced in PAMmer (Nelles et al. 2016). The RNA targeting using labeled Cas9 as CASFISH and CRISPRainbow can be used for the imaging (Deng et al. 2015, Ma et al. 2016); however, this system is not yet efficient to track single molecule. However, advancements like the use of MS2 tagging system where the stem-loop structure formed with the sequence binds to MS2 bacteriophage coat protein improves detection limit for the monitoring RNA in living cell (Han et al. 2020, Spille et al., 2019). Using Cas9-based approaches, multiple MS2 sequences can be inserted into the untranslated region of any gene so that after transcription fluorescent proteins or different dyes labeled with the coat protein can detect the transcribed RNA (Han et al. 2020, Spille et al. 2019). This method helps for the imaging and quantification of transcription activity of any gene.

Localization of Nucleolus Damages

The nucleolus is the most evident and discriminable nuclear subcompartment. It is a site of ribosome synthesis, a process that is highly synchronized to get proper cellular proliferation and cell growth. It can also intermingle with coiled (Cajal) bodies and participate in the maturation of non-ribosomal RNA. Studies revealed that it also has non-ribosomal functions (Boulon et al. 2010). The nucleolus can perceive stress and can play a significant role in synchronizing stress responses (Boulon et al. 2010). Several studies have reported that the nucleolus can be the target of toxic metal by using silver staining (Zou et al. 2012). Al treatment promoted expelling nucleolar material from the nucleus to the cytoplasm, as observed using silver staining (Shimizu et al. 2001). Medina et al. (1995) suggested Ag-NOR (silver staining of the nucleolar organizer) technique as a handy tool to study morphology and molecular properties of the nucleolus in onion root meristematic cells. Ag-NOR staining displayed brown color with the nucleolar region (Medina et al. 1995). Safranin is also a good nucleolus marker and is usually counterstained with aniline blue, Light Green or Fast Green. Different Safranin staining methods have been reported to give rise to specific color with nuclear materials, such as Safranin-Fast Green stain used to investigate chromosome and nuclei by staining them in red color and tannic acid ferric chloride-Safranin displaying red color with chromosome and nuclei (Ruifrok and Johnston 2001). Toluidine blue O gives purple-greenish blue color upon reaction with nucleic acids (Baker 1945, Ueda et al. 2011; Fig. 3I). Cytochemical stains available for the detection of DNA are Feulgen, acridine orange, ethidium bromide and DAPI, and for DNA/RNA methyl green-pyronin and azure B (Dashek 2000). The DNA-specific fluorochrome DAPI also helps in revealing the nuclear material (Zlobin et al. 2017).

Cell Death Markers

The study of plant cell death is relevant in stress physiology since it is often observed in response to exogenous stresses (Van Breusegem and Dat 2006). Cell viability can be assessed via recording the absorbance at 600 nm of Evans blue released from the stained tissue or by analyzing the blue-stained spots using the image processing program, such as ImageJ (Romero-Puertas et al. 2004, Ji et al. 2013). Fluorescein diacetate (FDA) and propidium iodide (PI) staining have also been widely used to analyze the cell death in plant tissues (San Miguel et al. 2012).

Apoptosis is measured by a terminal deoxynucleotidyl transferase (TdT), which labels the 3′-OH ends of DNA produced by DNA nicking, with biotin-conjugated dUTP (Gavrieli et al. 1992, Gorczyca et al. 1993). The TdT-mediated dUTP-biotin nick-end labeling assay associated with DAPI recognizes the nuclei that are or are not fated for elimination (Havel and Durzan 1996, Reape et al. 2008). FDA gives fluorescence upon reacting with living cell, while dead cells give no fluorescence. FDA is also used in combination with morphological analyses to assess the mode of cell death (i.e. to distinguish PCD from necrosis). The lack of fluorescence along with cytoplasmic condensation denotes that the cell has undergone PCD. The absence of fluorescence together with a lack of cytoplasmic retraction means that the cell was subjected to necrosis (Burbridge et al. 2006). One mechanism also consistently applied in the study of apoptosis is the CASPASE activity (cysteine proteases), typically caspase-3, which can be analyzed by using fluorogenic substrates (Palavan-Unsal et al. 2005).

Localization of Plant Secondary Metabolites

Plant secondary metabolism is affected by exogenous stresses, both abiotic and biotic. A clear example is the biosynthesis of phenolics that scavenge ROS produced upon stress (Berni et al. 2018). Upon exogenous stresses, plants react via changes in the secondary metabolism with a clear example provided by the phenylpropanoid pathway generating the building blocks of lignin/lignans, as well as flavonoids. Histochemistry can provide valuable support to downstream the chemical analyses of secondary metabolism in plants, by localizing, at the cellular/tissue level, the sites of production/accumulation of secondary metabolites. In this paragraph, we will provide examples of histochemical techniques applied for the visualization of specific classes of plant secondary metabolites.

Metabolites, such as polyphenols, flavonoids, anthocyanins and tannins, are the most studied plant molecules in recent years (Berni et al. 2019). Their particular chemical structure determines their involvement in many physiological plant processes (e.g. signaling and defense molecules, pigments) and they are instrumental in medical and pharmaceutical research for their antioxidant capacity (Berni et al. 2019). In this context, many studies are focused on the detection of plant metabolites, mainly in fruits, useful for their functional effects that increase the immune defenses through the diet (Shahidi and Ambigaipalan 2015). Biochemical studies are the most used methods for the detection of these molecules in plants, and they are often supported by molecular methods (Ignat et al. 2011). However, these techniques sometimes require elaborate sample preparation and particular extraction procedures that need to be adapted to the specific classes of molecules (Azmir et al. 2013). Moreover, in molecular/biochemical studies, it is impossible to establish the exact sites of metabolite localization. For these reasons, histochemical methods represent a complementary method to biochemical and molecular approaches that can help in the analysis in a relatively fast and accurate manner. Giuliani et al. (2018) described currently used fast histochemical colorations that allow metabolite localization in different plant tissues. For instance, the authors used the FeCl₃ staining procedure to detect the phenolic compounds, AlCl₃ for flavonoids and vanillin HCl for dark pigments. Ferreira et al. (2017) successfully used these staining methods for the detection of polyphenols, flavonoids and lignins in sections of Miconia albicans stem through light microscopy. Muravnik et al. (2016) used several different reagents in the glandular trichomes of Tussilago farfara. More specifically, the authors used the Wilson reagent (citric acid–boric acid 5/5, w/w) in absolute methanol to identify flavonoids, Toluidine blue O for phenols and the Nadi reagent (N, N-dimethyl-p-phenylenediamine) for terpenes, highlighting their specificity in secretory cells. The autofluorescence of some metabolites, such as phenols and flavonoids, can be used to localize these compounds at the cellular and subcellular levels in vivo by fluorescence or confocal microscopy (Roshchina et al. 2017, Talbi et al. 2020). These results underline the suitability of these methods as supporting tools for chemical and chromatography assays.

Metabolites as secondary compounds are produced through biochemical pathways involving the intervention of several enzymes. In this respect, we believe it interesting to mention, in this paragraph devoted to secondary metabolites, immunohistochemical techniques to localize specific proteins involved in the biosynthesis of secondary metabolites. The immunological reaction between the antibody and the target antigen ensures optimal resolution. For example, de Ávila et al. (2019) used antibodies for the localization of the enzymes polyphenol oxidase, peroxidase (POD) and stilbene synthase in Vitis vinifera L. berries using the epifluorescence microscope.

Localization of Lignin Deposition

Lignin is an aromatic macromolecule conferring mechanical support to plant cell walls (Christensen et al. 2001, Coleman et al. 2008, Polo et al. 2020). Lignin is particularly abundant in the secondary cell walls of cells composing the plant vascular tissue (Tobimatsu et al. 2013, Polo et al. 2020). Reinforcement of the cell wall by increasing lignin crosslinking may contribute to the protective cell mechanism against high metal concentration, acting as a sequestering system. Acidic phloroglucinol is commonly used for the histochemical analysis of lignin, giving a pink color (Sato et al. 2009, Zelko et al. 2012; Fig. 4B). Other probes include safranin-fast green and tannic acid ferric chloride-safranin, detecting lignin-dependent deposits with red colour (Ruifrok and Johnston 2001). Toluidine blue O reacts with lignin giving a blue-green to turquoise and phloroglucinol-HCl stains lignin with a red-violet color (Jensen 1962, O'Brien and McCully 1981). Lignin composition can also be studied, thanks to the different specificity of the dyes. For example, the Mäule staining, based on the sequential reaction of potassium permanganate-HCl-NaOH, was recently used in Cannabis sativa to differentiate lignin-containing S (syringyl) units from G (guaiacyl)-rich lignin: the S units give red color and the G units give brown staining (Behr et al. 2018). Iodine-sulfuric acid stains lignified walls in yellow (Baker 1945). The study of lignin using phloroglucinol staining on three species of Viburnum Linn., i.e. Viburnum punctatum, Viburnum coriaceum and Viburnum erubescens, showed the deposition of lignin in different regions of leaf, stem and roots, with the xylem as the chief lignin deposition place (Prabhu et al. 2011).

A method using fluorescence-tagged monolignols was developed to visualize lignin (Tobimatsu et al. 2013); sinapyl and p-coumaryl alcohols tagged with dimethylaminocoumarin and nitrobenzofuran were synthesized and fed to thale cress via transpiration stream. Such an approach allows us to visualize the dynamics of lignin formation in vivo and distinguish it from the preexisting lignin.

Localization of Oxidative Stress Markers

Oxidative stress in plants occurs due to excess ROS production indicated by an increase in superoxide anion and hydrogen peroxide. However, ROS significantly acts in plant cell signaling and plays its role under different stress circumstances. Moreover, excess ROS production can take place in different cellular compartments, such as chloroplasts, mitochondria, peroxisomes, plasma membrane, endoplasmic reticulum and nuclei (Mittler 2002).

In the plant cell, ROS accumulation can be imaged in vivo and in vitro by different histochemical methods (Kombrink and Schmelzer 2001, Sandalio et al. 2008) The generation of H₂O₂ in leaves and roots can be visualized by using the stain DAB (Fig. 4D). This compound was used for the first time by Graham and Karnovsky (1966) for the detection of POD by electron microscopy. DAB reacts with H₂O₂ in the presence of ROS producing a brown precipitate (Graham and Karnovsky 1966). This stain has been successfully used to analyze H₂O₂ accumulation in leaves and roots from different plant species exposed to heavy metals (Romero-Puertas et al. 2004, Rodríguez-Serrano et al. 2006, Rodríguez-Serrano et al. 2009; Table 3). As negative controls, sodium pyruvate or ascorbate can be used as H₂O₂ scavengers. To better image, brown H₂O₂-dependent spots in green tissues, the extraction of pigments can be done by boiling the tissue in ethanol (Romero-Puertas et al. 2004). In fresh tissues, H₂O₂ can also be imaged in vivo by using 2,7-dichlorofluorescein diacetate (H2DCFDA) under the confocal microscope (Ortega-Villasante et al. 2005, Sandalio et al. 2008, Rodríguez-Serrano et al. 2009, Pérez Chaca et al. 2014, Kováčik and Babula 2017; Fig. 4F). However, according to various studies, H2DCFDA reacts with other peroxides, such as hydroperoxides in the presence of PODs. So this dye should be considered a qualitative marker of oxidative stress rather than a specific indicator of H₂O₂ production (Sandalio et al. 2008). However, using specific H₂O₂ scavengers, such as sodium pyruvate or ascorbate, or with known H₂O₂ accumulation, it is possible to identify the presence of H₂O₂ in H2DCFDA fluorescence (Sandalio et al. 2008, Rodríguez-Serrano et al. 2009). This technique has successfully analyzed the differential H₂O₂/peroxide accumulation along with metal treatment in different types of cells in roots and leaves from several plant species (Ortega-Villasante et al. 2005, Rodríguez Serrano et al. 2006, Rodríguez-Serrano et al. 2009, Pérez Chaca et al. 2014). Imaging of H₂O₂ accumulation at a subcellular level can be carried out by using cerium chloride under electron microscopy, which allows discriminating the organelles involved in H₂O₂ production under different stress conditions, including metal toxicity (Romero-Puertas et al. 2004). This approach allowed to demonstrate three different waves of ROS induced by Cd in tobacco cell cultures (Olmos et al. 2003, Garnier et al. 2006). The use of specific inhibitors for various sources of ROS, such as DPI for NADPH oxidase and sodium azide for PODs, allows identifying the primary source of ROS for both in vivo and in vitro imaging (Sandalio et al. 2008).

A new noninvasive technique to monitor in vivo ROS accumulation is the use of genetically encoded ROS-sensitive fluorescent proteins, which allow subcellular ROS monitoring. One of these probes is HyPer, composed of cpYFP inserted into the regulatory domain of Escherichia coli OxyR. HyPer2 has two excitation spectrum peaks (405 and 488 nm, reduced and oxidized forms, respectively) and a single emission spectrum peak at 516 nm, which allow ratiometric analyses of H₂O₂ by the ratio 488/405 (Belousov et al. 2006). Hyper has been successfully used to analyze the effect of high light on H₂O₂ accumulation in chloroplasts and nuclei of tobacco leaves (Exposito-Rodriguez et al. 2017) and the effect of Cd on peroxisomal H₂O₂ production in Arabidopsis plants (Costa et al. 2010, Calero-Muñoz et al. 2019).

The accumulation of superoxide anion (⁠${\mathrm{O}}_2^{\bullet -}$⁠) in plant organs, such as leaves and roots, can be detected in situ by using nitro blue tetrazolium (NBT), which is reduced by the superoxide anion producing an insoluble blue-colored formazan complex (Table 3). To improve the visibility of superoxide-dependent spots in leaves, bleaching by immersion in boiling ethanol is recommended. Negative controls using tetramethylpiperidinooxy, a ${\mathrm{O}}_2^{\bullet -}$ scavenger, is recommended (Romero-Puertas et al. 2004).

Dihydroethidium (DHE) is known as a specific dye for ${\mathrm{O}}_2^{\bullet -}$ detection. DHE is the reduced form of DNA dye ethidium bromide and reacts with superoxide anions giving rise to the red fluorescent product 2-hydroxyethidium (Kalyanaraman et al. 2012; Table 3). DHE is used as a ${\mathrm{O}}_2^{\bullet -}$ marker in cell cultures and, according to the reported studies, in roots of pea plants treated with Al (Yamamoto et al. 2007), lupine roots exposed to Cd (Kopyra and Gwozdz 2003), soybean roots and pea root and leaf exposed to Cd (Rodríguez Serrano et al. 2006, Pérez Chaca et al. 2014), among others (Table 3).

MitoSOX Red is a cationic derivative of DHE, which specifically measures superoxide radicals in the mitochondrial matrix. It reacts with superoxide anion faster than DHE and forms the same product 2-hydroxyethidium (Zielonka et al. 2008). Imaging of ${\mathrm{O}}_2^{\bullet -}$ accumulation at the subcellular level can be carried out by using the method used to localize ${\mathrm{O}}_2^{\bullet -}$ radicals, which depends on the DAB precipitate in the presence of Mn²⁺ ions and ${\mathrm{O}}_2^{\bullet -}$ (Steinbeck et al. 1993). This technique is mainly based on the oxidation by ${\mathrm{O}}_2^{\bullet -}$ of Mn²⁺ to Mn³⁺, which further oxidizes DAB to an insoluble osmiophilic polymer (Steinbeck et al. 1993). This method has been used to analyze superoxide accumulation induced by Cd in pea leaves, showing accumulation in tonoplasts and plasma membrane (Romero-Puertas et al. 2004).

Nitric oxide is a small hydrophilic molecule, and being a secondary messenger in plant cells, it plays an essential role in several physiological processes like seed dormancy, seed germination, plant growth, flowering, stomatal closure, gravitropism, senescence and responses to different stresses (Neill et al. 2007). However, the time and the place where NO is produced is somewhat debatable, but to recognize the role of NO in physiological processes, it is imperative to determine its in vivo concentration in plant tissues. The fluorescent probe 4,5-diaminofluorescein-2 (DAF-2) diacetate has been generally used for the localization of NO in living tissues. The diacetate group is hydrolyzed by the cellular esterases, and DAF-2 reacts with NO and produces a highly fluorescent DAF-2 triazole, which is observed under excitation and emission at 495 and 515 nm, respectively (Sandalio et al. 2008).

Pitfalls and Solutions in Plant Histochemistry

Staining is defined as a process of color reaction that reveals the actual chemical constituents of the cells with the help of different dyes, stains, fluorochromes and microscopes (Spence 2001). However, for the detection of specific cell component, tissues and element, respective high affinity-specific dye is generally used (Hawes and Satiat-Jeunemaitre 2001). Therefore, histological detection through the use of specific stains gives a color product based on its chemical reaction with a specific element. Histochemical methods are essential for in vivo detection. However, in addition to this, these methods have a number of pitfalls in the quantitative analysis. It is evident that the accuracy of histochemical detection depends on image analysis and various fixation procedures of the plant specimens. Despite the complexities of quantitative analysis, histochemistry gives more direct information in addition to biochemical analyses and also provides spatial information. However, during histochemical staining, several problems can be encountered, which give ambiguous results, for instance:

• The precise localization of particular metal from metal complexes becomes difficult, because of the reactions between biomolecules and metals. Therefore, histochemical staining for single elements is sometimes unreliable.

• The selection of reagent is another problem, as at sometimes a reagent can show affinity toward many other elements. For instance, sodium rhodizonate has the highest affinity with Sr, but it is also used in the detection of Pb. Experimentally, it has been proved that Sr has more affinity toward sodium rhodizonate than Pb (Seregin and Kozhevnikova 2011).

• In some cases, the result gets distorted because of the errors during sample preparation. For instance, the use of ethanol with Ni-specific stain DMG causes crystal formation, which results in Ni redistribution in the plant cells (Mizuno et al. 2003). Moreover, nonspecific dyes can denature the reactive chemical groups or change non-reactive groups into reactive ones. One of the important assumptions for histochemical analysis is that the product formed by the staining can diffuse to other adjacent cells from its original location (Gahan 1984).

• Mishandling of plant tissue sections becomes the reason for less reliable result. The problem emerges at different stages of sample preparation, i.e. during cross-sectioning, fixing or infusion with resin, cells get broken or injured. This is the case for Ni, Zn and K, which are transported in cells through the symplast and are highly mobile inside plant cells (Krishnan et al. 2001, Seregin and Kozhevnikova 2011).

• Dye diffusion in plant cell is another pitfall that occurs during the staining process. Therefore, this can be escaped by applying the stain directly on the specific portion of the plant part subjected to the study. This results in washing out metal ions during the fixation and dehydration process. For instance, in most of the cases, Fe bound to ligand washes out during the staining procedures (Roschzttardtz et al. 2011). Therefore, to avoid such condition, one should carefully apply the dye for better binding with its specific element.

• For the imaging of cell viability, application of exogenous enzyme or probe is required. As the impermeable nature of plasma membrane is an obstacle for the detection of dead cells, therefore, prior to the enzymatic and ligand reaction, plasma membrane should be permeabilized. Furthermore, prior to the permeabilization, the tissue should be fixed in formaldehyde or glutaraldehyde, to reduce the chances of losing low molecular weight DNA (Palavan-Unsal 2005).

For a reliable outcome, it is important to compare the results obtained by histochemical analysis with other consistent methods, like electron microscopy, scanning electron microscopy, X-ray diffraction analysis and atomic absorption spectroscopy (Musumeci 2015). The sensitivity of histochemistry could be indeed a problem when low abundant compounds/elements need to be studied.

Conclusion and Future Perspectives

Histochemical methods are applied to detect subcellular components, secondary metabolites and cell wall components like cell wall polysaccharides and lignin, as well as for the localization of nucleic acids, promoter activity, minerals, metals, proteins, carbohydrates and lipids within cells and tissues. Their detection can be done on the basis of color-stain reaction using the microscope and photographic recording.

Histochemistry is a relatively easy, affordable technique widely used in several laboratories. It offers complementary information to high-throughput analytical methods for the analysis, e.g. plant mutants, the screening of experimental conditions impacting plant tissues and the choice of specific time-points along with kinetics for downstream detailed analyses. In addition, histochemistry is also used in other fields, namely plant taxonomy and medicinal botany. For example, histochemistry can be valuable for an initial assessment of the types of secondary metabolites produced in wild and ancient plant species (Berni et al. 2019). A histochemical study can be carried out to investigate the localization of specific secondary metabolites in different organs, as well as among different varieties within the same species. Such an approach could precede more time-consuming analyses requiring metabolite extraction and quantification.

Histochemical methods are especially valuable in the study of plant stress physiology, as it is possible not only to monitor the subcellular sites of secondary metabolite, enzyme and ROS production upon an exogenous stimulus but also to evaluate the impact on organelles. The possibility of synthesizing tagged building blocks used for the synthesis of biological macromolecules, such as lignin (see above), provides information on complex processes in vivo. Likewise, the use of click chemistry to specifically label cell components offers essential information on for example, the in vivo dynamics of plant membranes (Voiniciuc et al. 2018).

We, therefore, emphasize the importance of complementing high-throughput studies with histochemical analyses to get information on the spatial distribution of specific cell (macro) molecules, biogenic minerals and elements.

Acknowledgment

The authors would like to thank Head of the Department, University of Allahabad, Allahabad, India, for providing the necessary facilities to carry out the work.

Disclosures

Authors are declared that they don’t have any conflict of interest.

References