Tampion and Tampion (1987) defined an immobilized cell as a living cell that, by natural or artificial means, is prevented from moving freely from its original location to all parts of an aqueous phase of a system. On the other hand, the term “biofilm” comprises both organisms growing attached to a surface and organisms which grow as aggregates, where the cells are sticked together by extracellular polymeric substances (EPS) (Strieth et al., 2018). These are molecules produced by both algae and other microorganisms in response to physiological stresses encountered in the natural environment (Romaní et al., 2008). EPS serve to protect the cell from the surrounding environment and to bind the cell to a surface. EPS are composed of polysaccharides, proteins, nucleic acids, and phospholipids (Sheng et al., 2010).

Besides these recent definitions, many terms have been used in the literature by different research fields during the last decades. This section will give a glance of the extensive work on the topic. Two common approaches are used for increasing cell mass concentrations inside reactors: one is the use of a permeable membrane to retain cells; and the second one is using immobilized cell techniques. Membranes allow retention of the cells, while the liquid, substrate, and product out pass through the reactor. Thus, high cell concentrations can be achieved (Gao et al., 2015). However, the interest in the use of entrapped biomass is increasing because reactors with these types of biomass retention may offer high reaction rates, and solve some drawbacks that membrane methods have, as harvesting and maintenance (Qureshi et al., 2005).

Six different immobilization types have been proposed: covalent coupling, affinity immobilization, adsorption, confinement in liquid–liquid emulsion, entrapment in polymers and capture behind semi-permeable membranes (Mallick, 2002). These types of immobilization can be grouped into two groups: “passive and active,” using the natural ability of microorganisms to attach to (natural or synthetic) substrates or using flocculant agents, chemical attachment, and gel encapsulation, respectively (Moreno-Garrido, 2008). Other authors grouped them in three techniques, namely, “adsorption, entrapment, and covalent bond formation” (Qureshi et al., 2005). These two definitions are compatible because the techniques cited by Qureshi et al. (2005) can be included in the groups defined by Moreno-Garrido (2008) (Figure 2). Nevertheless, it should be considered that in some cases it may be difficult to classify a certain method as belonging to a specific group.

The entrapment and covalent bond formation techniques are active immobilization techniques. They require the use of chemicals, i.e., flocculant agents, chemical attachment or gel entrapment. This increases the cost of production and restricts further propagation or increase in cell concentration inside the reactor. With this technique, the biomass grows around the particles or surrounded by them and the size of the granules grows with time usually to several mm in diameter (de-Bashan and Bashan, 2010). The polymer is mixed with the microalgae cells and consequently stabilized with divalent ions to form immobilized microalgae beads through a nozzle (Lam and Lee, 2012). The density of the support particles is higher than the growing broth and bioparticles thus tend to remain in the lower section of the reactor. Since the immobilized beads are relatively large in size compared free cell cultures, a simple filtration method (e.g., sieving) is sufficient to separate the beads from water without significant energy input. Hence, handling of microalgae biomass is easy and feasible to be implemented at commercial scale (Lam and Lee, 2012; Castro-Ceseña et al., 2015). Some immobilization media, such as alginates, carrageenans and polacrylamide gel have been used for several purposes (Muñoz et al., 2009; de-Bashan and Bashan, 2010; He and Xue, 2010; He et al., 2014; Gagliano et al., 2017). Immobilization based on these polymeric matrices provides on low mechanical strength and can result in restrictive diffusion of substrate (Qureshi et al., 2005; Moreno-Garrido, 2008).

Many microalgae, bacteria, archaea, fungi, and protozoa cells have a natural tendency to attach (adsorb/adhere) to surfaces and grow on them (Huang et al., 2016). This technique, called adsorption, has been extensively used to attach microbial cells and is what was defined by Moreno-Garrido (2008) as “passive” immobilization. Normally, these processes are easily reversible. Adsorbent materials (carriers) for passive immobilization can be natural or synthetic (Ozkan and Berberoglu, 2013a; Gross et al., 2015; Katarzyna et al., 2015; Podola et al., 2017). The application of surface-attached growing algae is an interesting option, where algae are continuously harvested or extracted to obtain target compounds. Katarzyna et al. (2015) categorized this kind of immobilization as non-enclosure methods. Wimpenny et al. (2000) grouped in the term “aggregates” those accumulations of microorganisms that are developed on surfaces and those on interfaces were named as “biofilms.” Biofilms form most commonly at water/solid interfaces, although it can appear at an interface between two immiscible liquids like oil or water, at air-water interfaces and at gas-solid surfaces (Wimpenny et al., 2000). Similarly, a definition of a biofilm as an “aggregate of microorganisms,” in which cells are frequently embedded within a self-produced matrix of EPS adhere to a surface and/or to each other, binding firmly in the form of flocs and aggregates, property known as autoflocculation (Gonzalez-Fernandez et al., 2012) i.e., granular biofilms (González-Fernández and Ballesteros, 2012), was proposed by other authors with the aim to provide terminology useful and non-confusable between different research fields (Vert et al., 2012).

Algal biofilms can be established on any surface that receives sufficient light and moisture. These films are complex communities of both autotrophic and heterotrophic organisms (Callow, 2000). Thus, these films can contain different microorganisms, from diatoms and cyanobacteria to bacteria and/or protozoa. As an alternative, the term “photosynthetic biofilm” is often used to differentiate these films from those where the main source of energy is a dissolved or solid substrate (heterotrophic). However, the terms “algal biofilm,” “photosynthetic biofilm,” and “microalgal biofilm” have been used interchangeably to describe films in which microalgae constitute a significant portion of the microbial community (Irving and Allen, 2011).

Microalgal biofilms, composed of cyanobacteria and/or green microalgae, are ubiquitously distributed in almost all the photic aquatic environments. An important attribute of biofilms is that they both create and are functionally controlled by gradients of energy sources and chemical products (Bernstein et al., 2014). A determinant point involved in attached cultivation as defined before is the need of distinguishing two basic sub-types of “algal biofilm” (Wang et al., 2017). The first type is the biofilm consisting of pure and single species of microalga (axenic culture). For this biofilm, the attached cultivation is initiated from the concentrated algal biomass inoculum that is pasted onto the carrier material (Cheng et al., 2013; Liu et al., 2013; Shi et al., 2014; Zhang et al., 2015; Cheng et al., 2017). In this case, the biofilm diversity keeps stable during cultivation and no obvious contamination by exotic organisms occurs. The second type of biofilm is related with biofouling (Katarzyna et al., 2015). It may also start from axenic microalgal biomass, however, during the growth process, allochthonous bacteria, fungi and cyanobacterial also settle inside the biofilm and it becomes a diverse microbial community (Wang et al., 2017). Therefore, Mantzorou and Ververidis (2019) defined that a biofilm could be characterized as a consortium of microorganisms, embedded in EPS, forming a complex structure which is developed on solid surfaces.

Colonization of a surface is characterized by the alternation from freely moving cells in a, aquous medium, to a sessile, immobile community adhering to a surface. This transition is determinant on the microbial composition from the beginning of biofilm establishment to its maturity (Berner et al., 2015). The concentration of glycoproteins, proteins, cations and organic compounds at the surface can provide a nutritive zone for microorganisms compared to the bulk aqueous environment. Furthermore, fluid flow at the interface allows microorganisms to approach the hard surface. Once near the surface, microorganisms either get in contact with the surface by chemotaxis or by Brownian motion towards the surface in response to the chemical concentration gradient, and then the cell will form a temporary association with the microbes already present on the surface (Qureshi et al., 2005; Roeselers et al., 2007).

The initial adhesion of microbial cells to solid materials is mainly determined by the cell–solid and cell–cell interactions based on the physicochemical properties of the interacting surfaces, such as the thermodynamic balance of interaction energies (Sirmerova et al., 2013). Initially, cells can displace towards the conditioned surface in many ways: they can be transported by gravity, be transferred by advection or actively move via motile mechanisms (Ozkan and Berberoglu, 2013a). Attachment of most marine algae is thought to be facilitated by its specialized reproductive propagules which allow initial contact with a solid surface. Motile spores have the ability to respond to external stimuli such as surface chemistry and texture. On the other hand, non-motile spores are led by gravity and by the adhesive properties of the extracellular residual mucilage. After this initial contact all spores secrete adhesive materials which allow a permanent attachment to be made (Fletcher and Callow, 1992).

In freshwater, the primary colonizers are considered to be mainly diatoms, which are unicellular or colonial algae (Romaní et al., 2008). The spontaneous, often reversible, adsorption of organic and inorganic aqueous molecules to the solid surface is followed by a non-reversible secondary attachment of cells through the production of EPS. This provides protection from any toxic compounds that could harm the cells and from phagocytes and bacteriocides by granting a diffusive barrier to biofilm cells (Barranguet et al., 2005; Qureshi et al., 2005). The binding strength of EPS found in Algogenic Organic Matter (AOM) occurs by hydrogen bonding, electrostatic interactions and van der Waals forces between functional groups (Ozkan and Berberoglu, 2013a). Once the coating and the first biofilm layer is established, the colonization for other organisms is eased by the modification of the initial surface properties of the material (Lobelle and Cunliffe, 2011).

The second attachment, based on cell−surface interactions, has led to numerous adaptation strategies; e.g., surface charge (Hori and Matsumoto, 2010) and hydrophobicity of the cell walls and membranes can be adjusted by forming surface structures (Krasowska and Sigler, 2014), such as pili, curli, fimbriae (Bullitt and Makowski, 1995; Lasaro et al., 2009), and flagella (Guttenplan and Kearns, 2013) or by regulating EPS production (Romaní et al., 2008), all of which may improve adhesion to an habitable solid surface, establishing a stronger binding of the algal cells forming colonies (Rummel et al., 2017).

The micro-colonies produce more EPS, which serves to maintain the growing biofilm on the surface and increase its thickness during the development phase (Figure 3) as EPS supports cell–cell communication (Toyofuku et al., 2016). The biofilm can be colonized by secondary organisms and serve as a sink for other particulate matter in the environment (Wimpenny et al., 2000). The architecture of the biofilm develops during this phase in response to shear forces. Biofilms can form thick mushroom-like masses in low shear environments and may be flatter or form long strands in high shear environments (Costerton et al., 1995). This structure is shaped mainly by heterogeneous matrices with clusters made of cells and polymers, randomly distributed along a three-dimensional (3D) architecture and surrounded by water channels that sometimes extend from the bottom to the top of the biofilm (Lewandowski and Beyenal, 2003; Melo, 2005; Petroff et al., 2011; Toyofuku et al., 2016).

In this development phase, the cells living in the biofilm take up nutrients, mainly towards production of EPS rather than to biomass growth. When nutrients become scarce, cells must emigrate from the EPS matrix. Biofilm associated cells are able to release enzymes capable of breaking down the EPS matrix in times of nutrient starvation (Qureshi et al., 2005). Biofilm development is intrinsically linked to the accumulation of cellular and polymeric biomass. The loss of small or larger areas of biofilm biomass, called biofilm detachment, has been recognized as a part of the biofilm life cycle decades ago (Chang et al., 1991). In addition to abiotic growing factors that have an impact on the biofilm architecture, i.e., chemical factors such as cell detachment due to starvation or nutrient deficiency (Hunt et al., 2004; Wijeyekoon et al., 2004), and physical factors, i.e., abrasion, shear stress, sloughing and grazing (Stoodley et al., 2002; Telgmann et al., 2004; Celmer et al., 2008), there are also biotic factors as the microbial metabolic activity (Allison et al., 1998) and microbial gene expression (Kaplan et al., 2004) that influence biofilm detachment. A balance between attachment, growth and detachment of biofilm is important in the production and maintenance of a functional biofilm community (Garny et al., 2008; Woodcock and Sloan, 2017).

Several parameters affect how quickly biofilms form and mature. Microalgal adhesion to a hard surface is one of the main steps of biofilm development and strongly depends on different factors that can be grouped according to the origin/place where the effect occurs as: surface (interface), environmental and cellular factors (Qureshi et al., 2005; Irving and Allen, 2011; Liao et al., 2015). On other hand, other authors have divided these factors regarding their nature into: physicochemical properties, biological factors and environmental conditions (Barros et al., 2018). Both types of aggrupation can be exchangeable, as biological factors are mainly cellular and physiological properties that mostly occur at the interface. Important interface factors are the surface charge, surface energy, chemical nature, hydrophobicity (wettability), lubricity and surface topography. Environmental variables affecting photosynthetic biofilm formation include light availability, pH, hydrodynamic conditions, nutrient quality and quantity, temperature, carbon source and the target surface for attachment (Tan et al., 2020). Cellular influences are the microalgae genetics, growth stage and metabolic activity (Irving and Allen, 2011). Although there are many studies focusing on factors that impact the biofilm formation, especially surface physico-chemical properties of microorganisms and surface materials, studies comparing metabolism and experiments under in vivo conditions are uncommon or even absent for microalgae and cyanobacteria (Sekar et al., 2004a; Irving and Allen, 2011; Ozkan and Berberoglu, 2013a; Sirmerova et al., 2013).

The effect of substrate material on the algal attachment and attached growth is a complicated process (Schnurr and Allen, 2015). The physico-chemical properties of material surface such as hydrophobicity (Finlay et al., 2002.; Ozkan and Berberoglu, 2013b), surface energy (Genin et al., 2014), and dispersive surface energy (Cui and Yuan, 2013) are relevant parameters for the initial algal colonization, but this highly depends on the materials and model strain used.

From the cellular point of view, the genome, which determines the biofilm developmental potential of the organism, is one of the most important determinants of the biofilm structure. Thus, filamentous organisms might be expected to form a different multicellular array than non-motile coccoid organisms. Biofilms can be mainly regarded as approximately planar systems which include micro-colonies and more or less abundant EPS, which differentiates them from clearly filamentous structures (Wimpenny et al., 2000). Kokare et al. (2009) proposed that bacteria in biofilms express different phenotypic characteristics from planktonic counterparts. This is due to different genes transcribed in the planktonic and biofilm associated phases of the bacterial life cycle. Some genes may be expressed in response to a specific surface on which bacteria settle (Becker et al., 2001), e.g., genes involved in the attachment to chitin diverge from those required for attachment to abiotic, non-nutritive, surfaces such as glass or plastic (Keyhani and Roseman, 1996). Research on the role of metabolic and gene expression in microalgae biofilm formation is still a wide research field to explore, which can bring novel strategies to improve and enhance microalgae biofilm systems.

Considering the entrapped materials for biofilm formation, immobilized S. obliquus cells in polyvinyl and polyurethane were cultivated in order to remove nitrate from wastewater (Urrutia et al., 1995). Bench scale studies were performed using rotating disks of aluminum (Torpey et al., 1971) or polystyrene (Przytocka-Jusiak et al., 1984) to grow algae biofilms and reduce phosphorus and nitrogen levels in wastewater. The Algal Turf Scrubber (ATS) was developed by Adey and associates at the Smithsonian Institution (Washington D.C., United States) as a more natural means of maintaining water quality in model ecosystems (Adey et al., 1993). Plastic mesh has been frequently used to grow filamentous algae in a ATS by intermittently pouring water over the surface (Adey et al., 2011).

Some researchers have studied the differences in biofilm growth with different synthetic materials without any material property measurement, and differences in growth rates were found. Some representative studies of enclosed biofilm bioreactors screening carrier materials are for instance, the work of Sekar et al. (2004b) who used two species of green algae: Chlorella vulgaris and Nitzschia amphibian. Both green algae adhered most readily to titanium, stainless steel perspex and glass, while brass and copper showed fewer attached cells. Subsequent experiments with N. amphibian showed that cells were more likely to adhere to surfaces that had been roughened with sandpaper than the smooth control. Likewise, Johnson and Wen (2010) studied biofilm growth rates on six different carriers (Supplementary Table S1). They observed that polystyrene foam yielded significantly higher biomass productivities than the other materials tested. On the other hand, Genin et al. (2014) examined six materials for their capability of supporting algal growth in a parallel airlift bioreactor. In this case, cellulose acetate showed the highest biomass productivity. Shen et al. (2014) evaluated the microalgal biofilm formation on nine carrier materials and the higher biomass productivity was observed using glass fiber reinforced plastic.

Christenson and Sims (2012) tested eight synthetic and natural carrier materials for the ability to support algal attachment and found that cotton rope showed the best performance. Gross et al. (2013) screened 16 potential biofilm carriers and found that cotton duct was the best material for microalgal biofilm growth. Likewise, (Moreno Osorio et al., 2019b; Moreno Osorio et al., 2020) reported efficient attached microalgal growth of C. vulgaris ACUF_802 and ACUF_809 as well as to S. vacuolatus ACUF_053 on cotton fabric. Similarly, Chuah et al. (2020) found that attached growth of C. vulgaris on cotton duct served as a better supporting material than polystyrene foam as it promoted microalgae attachment at low-cost, supporting an easy microalgae harvesting process.

An extensive and detailed study was performed by Gross et al. (2016), 33 synthetic materials with smooth surface were tested. The materials were grouped as metals, plastics and rubbers to evaluate their surface physico-chemical properties. Similarly, Barros et al. (2018) studied the influence of physicochemical properties of selected microalgae strains, growth medium composition and six carrier materials, using a lab scale methodology that allowed to evaluate multivariables in parallel.

Other authors used the same type of bioreactor configuration when testing different substrate materials. Naumann et al. (2013) developed a solid-state bioreactor, denominated “twin layer system” that combines a nutrient substrate and works as a support layer of an attachment surface where the biofilm grows. They evaluated the biofilm formation performance of four microalgae strains on plain printed paper. They concluded that Phaedactylum tricornutum has the higher biomass productivity (Supplementary Table S1). Complementary, (Shi et al., 2014) tested nylon filter sheets as attachment substrate material for the biofilm formation of Hallochlorella rubescens.

Studies for testing materials with different surface textures have demonstrated that the patterns on the substrate cannot only offer larger areas for cell attachment, but also act as anchor points sheltering cells from shear stresses (Huang et al., 2018). As expected, higher biomass productivity is achieved on materials with relatively larger surface area. Cao et al. (2009) confirmed that micro-dimple surface features of corrosion resistant steel sheets could significantly enhance the attachment of algal cells to the carrier, compared to a surface without microdimple features.

Additional materials have been used at laboratory scale to screen the biofilm formation capability of microbial consortia or microalgae axenic cultures, most of them aiming for the selection of a cheap and easily produced surface material that could be applied at large-scale in-land systems. The system used by Ozkan et al. (2012) is notable for its use of concrete as a cultivation surface, which can mean a significant reduction in the maintenance cost. The same material was used by Sukacova et al. (2015) with effective results. Locally produced materials as glass (Schnurr et al., 2013), foam PVC with surfaces sanded in order to promote microbial attachment and polyvinyl chloride (Posadas et al., 2013) were selected based on availability, non-toxicity, ease of manipulation and costs.

Various types of natural materials have been used as carriers for the microalgal biofilm cultivation (Supplementary Table S1). The advantages of a natural carrier material are the low cost and the environmental sustainability of the biofilm system, especially compared to the immobilization technologies where (although these can employ natural polymers, i.e., alginate and carrageenan), the high cost of the polymeric matrix prohibits their use at large scale (Hoffmann, 1998; Christenson and Sims, 2011). Several immobilization media, such as alginates, carrageenans and polacrylamide gel have been used for entrapped the microbial biomass (Moreno-Garrido, 2008). Akhtar et al. (2003) and Akhtar et al. (2008) provided an immobilized microalgal biosorption process using a highly porous, ridged, physically strong and low cost immobilization matrix (luffa sponge) that consists of discs obtained from matured dried fruit of Luffa cylindrica for the removal of Ni(II) and Cr (III) using Chlorella sorokiniana as model organism. Surface immobilization of the microalgae on luffa sponge, providing direct contact of the biomass to the metal solution, was better suited for metal biosorption than microalgae cells enclosed or beaded immobilized in a polymeric gel structure.

Filter paper was used as carrier material for algal growth vertically, and differences in biomass productivity were observed between three microalgae biofilms (4.2–6.1 m⁻² d⁻¹) (Naumann et al., 2013). Zhang et al. (2017) explored the feasibility of using lignocellulosic materials as biofilm carriers, which had a similar chemical composition to the algal cell wall. Of the materials tested, pine sawdust as carrier achieved high biomass productivity (10.2 g m⁻² d⁻¹). Lignocellulosic-based materials are generally characterized by a high surface energy and achieved greater attachment than synthetic polymers which have a low surface energy (Christenson and Sims, 2012). Moreover, lignocellulosic materials are hydrophilic natural polymers and have a good liquid-holding capacity (Zhang et al., 2017). The biofilm productivity was found to be increased with the increase of material surface roughness (p < 0.05) (Cao et al., 2009). Fica and Sims (2016) characterized the biofilm formation and nutrient removal of a microbial consortium dominated by algae species. The biofilm was grown on cotton cords in a Rotating Algal Biofilm Reactors (RABR) as described by Christenson and Sims (2012). They found that different concentrations of total organic carbon and temperature influenced the biomass productivity.

The rotating algal biofilm system proposed by Gross et al. (2013) was a conveyor belt system that had cotton as carrier material. The carrier moved around several rotating shafts, passing through a reservoir of nutrient-rich liquid at the bottom. Cotton duct promoted a high biofilm productivity in other studies with different alga strains (Christenson and Sims, 2012; Gross et al., 2016; Zhuang et al., 2016; Carbone et al., 2017). A clinoptilolite rotating photobioreactor (cRPB) was used by Young (2011) for removal of nitrate and ammonium from synthetic wastewater, the zeolite surface can be bio-regenerated through continued nitrogen removal. A microbial consortia biofilm productivity of 12 m⁻² d⁻¹ was obtained. Two component biodegradable carriers for Biofilm Airlift Suspension (BAS) reactors were used to investigate the oxygen transport inside the biofilm and the development of the biofilm structure (Hille et al., 2009). The carriers were composed of PHB (polyhydroxybutyrate), which is easily degradable, and PCL (caprolactone), which is less easily degradable by heterotrophic microorganisms (Hille et al., 2009).

Talukder et al. (2014) studied for the first time the immobilization of microalgae on exogenous fungal mycelium. Mycelium is a natural carrier material that can be obtained free or at low-cost as waste from fungal fermentation processes that produce valuable products, i.e., enzymes or soy sauce. The positively charged hyphae interact with the negatively charged algal cell surface and cause flocculation (Alam et al., 2016). The occurrence of filamentous microorganisms could thus be considered as a beneficial factor for microalgal biofilm cultivation by the promotion of algal cell attachment on the substrate layer. Meanwhile, respiration of the microorganisms could enhance algal productivity by decreasing O₂ inhibition of algal growth (Bai et al., 2015).

The simple identification of the microbial groups is not informative enough, as the microbial activity is influenced by the biofilm spatial structure (Barranguet et al., 2004). Many different methods to characterize the properties of biofilms including detection, imaging, and sensing methods have been proposed in the literature (Neu and Lawrence, 2015). Sensing and imaging differ from each other in that sensing refers to the quantifiable measurement of some analyte or property of the subject or system, while imaging is used to generate visual information (topography, area, colour) related to a subject, and some combine it with the detection of analytes or even concentration measurements (Saccomano et al., 2021).

Techniques that do not disrupt biofilm structure and its microgradients have been given priority for understanding biofilm formation and development (de Beer et al., 1993; Zippel and Neu, 2005). Non-destructive methods for examining photosynthetic biofilms (Table 3) include various forms of microscopy and chemical imaging: simple light microscopy, Helium Ion Microscopy (HIM) (Schmidt et al., 2021); Scanning Electron Microscopy with Energy Dispersive X-ray analysis (SEM-EDX) (Moreno Osorio et al., 2019a), Confocal Laser Scanning Microscopy (CLSM) (Neu and Lawrence, 2014; Moreno Osorio et al., 2020), Confocal Raman Microscopy (CRM¹) (Moudříková et al., 2017; Keleştemur et al., 2018), Pulse Amplitude Modulation (PAM) (Wimpenny et al., 2000; Moreno Osorio et al., 2020), Vibrational Spectroscopy (ATR-FTIR spectroscopy) (Fanesi et al., 2019), Time-of-Flight Secondary Ion Mass Spectrometry (ToF-SIMS) (Benettoni et al., 2019) and imaging analysis (De Muynck et al., 2009).

An increasing trend in nanoscale research is the combination of more than two techniques (high resolution microscopy + chemical imaging) to analyze different parameters in parallel to greatly improving the sensitivity and spatial resolution to visualize and quantify nutrients, metabolites and toxic elements in single cells at the subcellular and cell to cell level. Many of these are considered emerging imaging technologies that can be combined to correlate elemental, isotopic and molecular information from a region of the cell or from the entire cell in two or three dimensions (Decelle et al., 2020).

Optical methods have arisen as promising techniques in biofilm studies. Although biofilms can be observed with an unaided eye, imaging their structure, distribution of EPS and microbial community structure requires the use of several types of microscopy instruments combined with various probes, such as fluorescent proteins (FPs) used as reporter genes and labels or fluorescent in situ hybridization (FISH) probes (Azeredo et al., 2017).

HIM resembles a field-emission scanning electron microscope (FE-SEM) in many aspects, but several advantages are provided by the use of helium ions rather than electrons, including higher surface sensitivity, larger depth of field and a straightforward charge-compensating electron flood gun, which enables imaging of non-conductive samples, rendering HIM a promising high-resolution imaging technique for biological samples. The key remarkable capabilities of HIM in bio-imaging, in terms of spatial resolution, very large depth of field, and the ability to image non-conductive samples as biofilms without the application of conductive coatings support the growth in the number of studies which has been published in related fields and the opportunity for further developments (Schmidt et al., 2021). However, every problem in microscopy cannot be solved by HIM alone. In subcellular chemical imaging, for instance, correlative workflows combining nanoanalytics (e.g., X-ray spectroscopy and SIMS) and high-resolution electron microscopy are currently being established (Decelle et al., 2020).

In ToF-SIMS, a release of atoms from the outer few nanometers of the surface is caused by the bombarding with ions of the sample surface, and their identification is based on their mass. In SIMS imaging, high resolution in the mass spectrum comes at the price of lower image spatial resolution. However, development of the Nanoscale Secondary Ion Mass Spectrometry (NanoSIMS) allowed imaging at sub-micron resolution while maintaining high mass resolution (Azeredo et al., 2017). LeTourneau et al. (2018) used SEM, HIM, and NanoSIMS as correlate imaging tools to show that phenazine-1-carboxylic acid, produced by rhizobacteria in unirrigated wheat fields, and soil moisture promotes biofilm formation at root surfaces. The stable isotope labelling in combination with NanoSIMS, electron microscopy and HIM was used to study the carbon and nitrogen cycles. SIMS has demonstrated to be a powerful tool to study the unicellular and cell to cell interaction of microalgae (de Bashan et al., 2016) that make it a promising technique for the characterization of nanoparticles in biological media. In fact, high spatial resolution, its sensitivity and mass resolution offer label-free chemical 3D imaging capabilities down to the nanometer scale. Benettoni et al. (2019) used ToF-SIMS in an experiment on C. vulgaris biofilms to confirm the presence of TiO₂ nanoparticles and to clarify their 3D distribution within this complex system. Similarly, Moreno Osorio et al. (2019) performed ToF-SIMS analyses on biofilm samples to analyze the distribution of phosphorus-containing molecular species in Chlorella pyrenoidosa and C. vulgaris biofilms. Furthermore, the high resolution and the wide range of molecular species that can be 3D-detected with this imaging technique allowed to understand the role of the cotton fabric as algal biofilm support material and confirmed that the cotton carrier does not accumulate phosphates and thus phosphate removal was the result of biofilm metabolic activity.

The favorite types of microscopies among biofilm researchers are those that allow the examination of living and fully hydrated biofilms. In addition, sophisticated image acquisition devices that can selectively image and stimulate various probes are often required when more than one type of multicolored probe is used simultaneously (Lewandowski and Boltz, 2011). Among them, CLSM enables the in vivo reconstitution of the 3D structure of microbial biofilms in their naturally hydrated form (Zippel and Neu, 2005). CLSM provides simultaneous identification of the different components and the information about the 3D structure of the biofilms, either by using specific fluorescent dyes for bacterial DNA or EPS glycoconjugates or by algal autofluorescence (Staudt et al., 2004; Neu et al., 2010; Neu and Lawrence, 2015).

Most of the understanding about biofilm architecture and composition (Chen et al., 2007; Neu and Lawrence, 2014) as well as their relation to mass transfer (Yawata et al., 2010) and metabolism is gained from rather simple CLSM point scanners. Several studies have been carried out using CLSM as informative technique for microalgae biofilm formation performance (Norton et al., 1998; Barranguet et al., 2004; Mueller et al., 2006; Blanken et al., 2014; Cole et al., 2014; Sommerfeld Ross et al., 2014; Zhang et al., 2017). A set of computational tools for extraction of the biofilm morphology from the 3D structural data obtained from CLSM imaging, and the accompanying web-based interface was described by Xavier et al. (2003).

A variation of the reflection microscopy technique employing a CLSM is defined as confocal reflection microscopy (CRM²). This technique allows the visualization of 3D samples without fluorescent labeling and fixation. Yawata et al. (2010) achieved a successful non-destructive monitoring of the entire biofilm development of Streptococcus mutans, on microfluidic-device experiments by a modified CRM² technique. This modified version of CRM² with continuous gain adjustment was termed Continuous Optimizing CRM (COCRM) and the mean biovolume measured showed good correlation with those calculated based on SYTO9 staining.

Biofilms are of outstanding interest for biological wastewater treatment, however in situ and real time techniques for analyses of biofilm systems (aerobic and anaerobic) are still limited. In contrast to invasive techniques for imaging analysis, such as FISH, Patzold et al. (2006) were able to identify anaerobically ammonium-oxidising (anammox) bacteria without pretreatment in samples just by its Raman vibrational signature. The presented results provided new insights into the complex interactions of different organisms in microbial communities without interfering with them. On the other hand, analytical techniques used to investigate the presence of lipids in the selection of microalgae for the optimal production of biofuels include Gas Chromatography–mass spectrometry - GCMS (Bigelow et al., 2011), Liquid chromatography–mass spectrometry - LCMS (MacDougall et al., 2011), Nuclear magnetic resonance – NMR (Nuzzo et al., 2013), Fourier-Transform Infrared Spectroscopy - FTIR (Dean et al., 2010), and Raman spectroscopy (Wei et al., 2014). Confocal Raman spectroscopy on the other hand, allows access to full spectral information with high spatial resolution (Sharma et al., 2015). Combination of Raman spectroscopy with confocal optical microscopy resulting in 3D spatial characterization of the samples has resulted in CRM¹ becoming the method of choice for label-free and real-time monitoring of various biological samples, including microalgae and bacterial biofilms (Murdock et al., 2008; Efeoglu and Culha, 2013; Wei et al., 2014; Moudříková et al., 2017; Keleştemur et al., 2018).

The pioneer study of Valladares Linares et al. (2016) assessed three novel in-situ biofouling characterization techniques on membrane systems: i) nuclear magnetic resonance (NMR), ii) oxygen imaging with planar optodes, and iii) optical coherence tomography (OCT). These non-destructive tools can elucidate the interaction of hydrodynamics and mass transport on biofilm accumulation in membrane systems. OCT is a key emerging imaging technique of the last decade in biofilm research. Remarkable features of this technique are the in-situ application, the real-time-acquisition of multi-dimensional datasets as well as the fact that no sample preparation is necessary allowing for a non-invasive and complete characterization of an unaltered biofilm structure (Wagner and Horn, 2017). A significant advantage of OCT against other microscopy imaging techniques is the availability of investigating biofilms directly inside the cultivation device (e.g., biofilm reactor) under operational conditions. 2D and 3D datasets contain a representative description of the overall biofilm structure (Wagner and Horn, 2017). In-situ characterization using NMR techniques were successfully applied for non-invasive online monitoring of biofilm development, sloughing, forced detachment, and chemical cleaning. It allows quantification and visualization of development of biofilm and the cell interactions with the surrounding fluid at the mesoscale (McLean et al., 2008; Wagner et al., 2010).

Planar optodes use luminescent O₂ indicators immobilized in an O₂ permeable polymeric matrix, which can be coated on glass or foils surfaces. The principle is based on measuring the dynamic collisional quenching of the indicator luminescence by O₂ (DeGraff and Demas, 2005). Optical O₂ sensing was first developed in the medical field for blood gas analysis and was introduced in aquatic microbiology when fiber-optic O₂ microsensors were developed and applied to biofilm studies (Kühl et al., 2007; Neu and Lawrence, 2015; Khosravi et al., 2020; Saccomano et al., 2021).

In microalgal biofilms, PAM fluorimetry allows the measurement of algal biomass on undisturbed biofilm samples, maximum quantum yield and algal photosynthetic activity (Genty et al., 1989; Barranguet and Kromkamp, 2000). Variable fluorescence emerged as a promising and non-intrusive tool to estimate photosynthesis in phytoplankton (Kolber et al., 1998). The principle of variable fluorescence is also used by PAM fluorimeters (Schreiber et al., 1995) and this technique has been used to measure algal photosynthesis, although mainly on laboratory cultures. Increasing applications of PAM fluorimetry for biofilm studies have been reported in the literature of the past decade (Eggert et al., 2006; Häubner et al., 2006; Malapascua et al., 2014; Manso et al., 2014; Ng et al., 2014; Schuurmans et al., 2015; Wang et al., 2016b).

De Muynck et al. (2009) evaluated the algal fouling on concrete by means of colorimetric and image analysis. The fouling intensity parameter and the image analysis is based upon the quantification of the area covered by algae by means of threshold analysis on the a* (green to red axis) and b* (blue to yellow axis) coordinates of the CIELab colour space of ImageJ software (Rasband, 1997–2008). This analysis allows a clear distinction between fouled and unfouled areas of concrete. Imaging analysis has been an effective tool to follow the biofilm growth in external facades and monuments (Sanmartin et al., 2010; Fernandez-Silva et al., 2011; 2011; 2012). Colorimetric measurements and image analysis were used to evaluate the intensity of the alga Klebsormidium flaccidum biofilm to evaluate the intrinsic properties of porosity, roughness, and surface pH on the susceptibility of mortars to biodegradation (Tran et al., 2012; Tran et al., 2014). Eyssautier-Chuine et al. (2016) and Carbone et al. (2017) combined the analysis of color changes in the carrier surface (stone and cotton duct, respectively) and the algal photosynthetic activity using PAM fluorometry. Both studies concluded that both techniques are complementary in the analysis of microalgae biofilm formation.

Another study that combined and correlated different non-destructive imaging techniques analyzed the early colonization stages of Chlorella biofilms by coupling imaging analysis techniques and mathematical methods (Moreno Osorio et al., 2020). Different parameters were measured correlating imaging techniques: the biomass growth and health condition were measured using PAM fluorometry; carrier colonization was studied by imaging analysis (Weka Segmentation) and through CIELAB colorimetry the green tones variations in the biofilm were measured. Biofilm vertical distribution and structural features were estimated by multichannel CLSM, and some parameters were aligned to the Gompertz model (Moreno Osorio et al., 2020). However, studies of this type are not common (Wagner and Horn, 2017; Fanesi et al., 2019), the combination of different techniques in bacterial biofilms is an extensive research field (Yang et al., 2013; Decelle et al., 2020) as has been shown along this section.