# Illuminating microalgal biofilms using advanced photonic imaging techniques

Julia Mougin, Anne-Sophie Pavaux, Julien Deschamps, Vlad Costache, Filipa Lopes, Romain Briandet

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Julia Mougin, Anne-Sophie Pavaux, Julien Deschamps, Vlad Costache, Filipa Lopes, et al. Illuminating microalgal biofilms using advanced photonic imaging techniques. Algal Research - Biomass, Biofuels and Bioproducts, 2025, pp.104018. 10.1016/j.algal.2025.104018. hal-05005963

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Illuminating microalgal biofilms using advanced photonic imaging techniques

Julia Mougin, Anne-Sophie Pavaux, Julien Deschamps, Vlad Costache, Filipa Lopes, Romain Briandet

<figure><img src="image_2.png" /></figure>

PII: S2211-9264(25)00127-4

DOI: https://doi.org/10.1016/j.algal.2025.104018

Reference: ALGAL 104018

To appear in: *Algal Research*

Received date: 20 September 2024

Revised date: 12 February 2025

Accepted date: 22 March 2025

Please cite this article as: J. Mougin, A.-S. Pavaux, J. Deschamps, et al., Illuminating microalgal biofilms using advanced photonic imaging techniques, *Algal Research* (2024), https://doi.org/10.1016/j.algal.2025.104018

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# Illuminating microalgal biofilms using advanced photonic imaging techniques

Julia Mougin<fnref n="1" />,<fnref n="2" />, Anne-Sophie Pavaux<fnref n="1" />,<fnref n="2" />, Julien Deschamps<fnref n="1" />,<fnref n="3" />, Vlad Costache<fnref n="3" />, Filipa Lopes<fnref n="2" />* & Romain Briandet<fnref n="1" />*

Co-authors' email: julia.mougin@ifremer.fr ; annesophie.pavaux@gmail.com ; julien.deschamps@inrae.fr ; vlad-costache@inrae.fr

**Keywords:** Photosynthetic biofilm, Microalgae, Bacteria, Multi-scale Imaging, Fluorescence microscopy

## 1. Abstract

In recent years, microalgal biofilms have garnered significant interest primarily due to their potential to transform wastewater treatment and overcome the limitations associated with conventional microalgal planktonic-based cultivation methods. Advances in imaging technologies have provided insight into the complex dynamics of microalgal biofilms, which could help in understanding both biofilm structure and function, and thus to optimize microalgae biofilm-based technologies. In particular, photonic microscopy allows researchers to study the architectural nuances, physiological responses, as well as spatial organization and interactions within biofilm communities. Despite these significant advances, there is still a lack of consistency in approaches and analytical techniques, which hinders standardized studies. Therefore, there is a need for common protocols and benchmarks in photosynthetic biofilm imaging. This review focuses primarily on key photonic imaging technologies, including confocal laser scanning microscopy (CLSM), light-sheet microscopy, histology coupled with a digital slide scanner, and optical coherence tomography (OCT). Principles, advantages/limitations and current applications of each technique in the context of microalgal biofilm studies, are presented allowing researchers to make reasoned decisions for their specific investigative endeavors.

## 2. Introduction

Photosynthetic biofilms, intricate communities of surface-attached microorganisms driven by light energy, play a significant role in aquatic ecosystems, contributing to ecological processes. The interplay of cyanobacteria, microalgae and other photosynthetic microorganisms orchestrates the

<footnote n="1">¹ Université Paris-Saclay, INRAE, AgroParisTech, Micalis Institute, 78350 Jouy-en-Josas, France</footnote>

<footnote n="2">² Université Paris-Saclay, CentraleSupélec, Laboratoire Génie des Procédés et Matériaux (LGPM), 91190 Gif-sur-Yvette, France</footnote>

<footnote n="3">³ Université Paris-Saclay, INRAE, AgroParisTech, Micalis Institute, MIMA2 Imaging Facility, Microscopie et Imagerie des Microorganismes, Animaux et Aliments, 78350 Jouy-en-Josas, France</footnote>

<footnote>* Corresponding authors: filipa.lopes@centralesupelec.fr & romain.briandet@inrae.fr</footnote>
conversion of light energy into chemical bonds, thereby shaping the availability of resources for an array of cohabiting organisms. While often characterized as a biofouling nuisance [1, 2], these biofilms exhibit a spectrum of promising applications ranging from environmental bioremediation to biotechnological innovation [3-5].

Notably, the interest in photosynthetic biofilm-based systems has increased in the past years [6-8]. Their capacity to effectively remove pollutants has led to a surge in research efforts aimed at harnessing microalgal biofilms, including photogranules, as a promising and sustainable solution in wastewater treatment applications worldwide [5, 7-12]. The potential of combined bio-product production further strengthens the overall sustainability and competitiveness of the system. In microalgae farming, biofilm-based cultivation can circumvent various limitations and drawbacks associated with the commonly used planktonic-based methods (open raceway cultures and closed photobioreactors), including but not limited to the high costs and energy consumption associated with harvesting and dewatering [13, 14]. In an era marked by a transition towards renewable energy and a circular economy to achieve sustainability, biofilm-based cultivation offers a pivotal alternative for the development of a resource-efficient industry and is garnering growing interest from researchers[14, 15].

Microalgal biofilms, whether mono- or multi-species, manifest a three-dimensional (3D) structure affixed to surfaces, with microorganisms embedded in an Extracellular Polymeric Substance (EPS) matrix [16]. This architecture (spatial arrangement of microorganisms, extracellular polymers and particulates) can contain a consortium of microorganisms including not only photosynthetic organisms but also non-photoautotrophic bacteria, fungi, archaea, and protozoa. These biofilms can give rise to heterogeneous cell clusters interconnected by channels forming a complex and dynamically regulated network governed by an inherent machinery system. Such microbial assemblages benefit from increased resistance to stress and division of labor, with physiological heterogeneity influenced by numerous biotic (including species, cell-cell signaling) and abiotic factors (including nutrient availability, light, hydrodynamics) [17, 18]. By deciphering the intricacies of these ecosystems, researchers can gain critical insights into biofilm behavior, thereby facilitating the optimization of biofilm-based culture systems and the prevention of undesirable biofouling [19].

As a result, recent years have witnessed a significant advances in imaging technologies, providing unprecedented capabilities to explore the complex community of photosynthetic biofilms [20-23]. Multimodal imaging tools have indeed enabled to investigate the dynamic mechanisms of development, reactivity, interaction between species and the surrounding environment, providing a better understanding of the structure/function/biodiversity relationships. However, there is still a lack of consensus among the diverse approaches and techniques, applied to the investigation of microalgal biofilms, limiting the number of standardized studies. This calls for a concerted effort to establish
common protocols and benchmarks to advance the field of microalgal biofilm technology. In this context, imaging techniques could help filling two major knowledge gaps: understanding interspecies interaction and the role of biofilm architecture on productivity and stability.

In the following sections of this review, we embark on a comprehensive exploration of the prominent photonic imaging technologies - Confocal Laser Scanning Microscopy (CLSM), Light-sheet microscopy, histology coupled with a digital slide scanner, and Optical Coherence Tomography (OCT) - that have emerged as potential tools in the study of microalgal biofilms. This review attempts to provide researchers with a comprehensive resource to navigate the landscape of imaging technologies in the field of microalgal biofilm research and in particular, their input to make the microalgae-biofilm technology progress. The Table 1 summarizes the advantages, drawbacks and main applications of each technique.

### 3. Bridging major knowledge gaps in microalgae biofilm technology

Though the interest of the photosynthetic biofilm-based technologies, their application at large scale is still scarce. More studies are therefore required to demonstrate their potential as highly productive, efficient, stable and robust technologies. In this context, a fully understanding of the relationship between biofilm structure/function/biodiversity is crucial and can be achieved by using imaging techniques.

Microalgae cultures are most often characterized by the presence of other microorganisms, either related to an external contamination [24] or by the supply of specific species that are known to improve algae performance [25]. In both situations, synergistic and/or antagonist relationships are established that must be comprehended for better exploitation and control of the cultivating systems [26]. As a matter of illustration, we can cite the particular case of a *Chlorella* sp. culture contaminated by a predatory obligate cyanobacteria, the *Vampirovibrio chlorellavorus*, that led to culture collapse [27]. It should be mentioned that while different contaminant organisms have been reported in planktonic cultures of photosynthetic organisms, including but not limited to bacteria, protozoa, microalgae, or fungi, information regarding microalgal biofilms is still scarce [28-31]. On the other hand, it has been demonstrated that the interaction between microalgae and other microorganisms, mainly bacteria, in biofilm-based cultures do improve biomass productivity [32] and nutrient removal in wastewater systems [33, 34].

Assessing the role of biofilm structure on process productivity and stability is another important scientific challenge to be addressed in the field. Indeed, within these structures, the chemical and light gradients over biofilm depth affect single cell behavior (physiology, activity) and finally determine the overall biofilm functioning, properties and productivity. Therefore, as soon as limitations occur, activity loss and sloughing-off occur which compromise the cultivation technology [35]. Therefore, it
is crucial to maintain an optimal biofilm thickness and to determine the optimal harvesting period to ensure structural integrity, process stability and maximize productivity [36]. On-line monitoring tools and comprehensive studies of biofilm dynamics are therefore essential for characterizing biofilm properties and gaining deeper insights into biofilm time-resolving architecture and productivity [37].

As such, while microalgal biofilms hold significant promise for large-scale cultivation, their full potential remains underutilized due to the limited understanding of the structural, functional, and biological interactions. An endeavour, that could be overcome thanks to imaging techniques.

# 4. Principles of fluorescence-based approaches

Considering that mostly used techniques are fluorescence-based, we will first discuss methods to selectively contrast photosynthetic from non-photosynthetic organisms and fluorescence markers to assess physiological, metabolic and chemical features in biofilms.

## 4.1. Contrasting photosynthetic cells

Chlorophyll autofluorescence is highly employed for monitoring microalgal growth in bioprocesses [38, 39]. In case of photonic imaging, it has been also widely used for the detection, *in situ* and non-destructively, of photosynthetic biofilms [20, 40, 41]. In microalgae chlorophyll is typically excited with wavelengths around 430-450 nm (blue light) or 640-660 nm (red light), with peak emission in the far-red region, around 680-700 nm [39]. Furthermore, along with microalgae, different photoautotroph organisms in particular cyanobacteria, purple bacteria or green sulfur bacteria, may harbor distinct fluorescence properties due to the presence of various photosynthetic pigments. For instance, cyanobacteria possess both chlorophyll and phycobilin pigments (phycoerythrin, phycocyanin, allophycocyanin), as well as scytonemin, an UV protecting compound resulting in a green autofluorescence [42, 43]. Cyanobacteria exhibit fluorescence peaks in the orange (phycoerythrin ~570 nm) and red regions (phycocyanin ~630 nm and allophycocyanin ~660 nm), which can overlap with chlorophyll signals. To optimize imaging and minimize overlap, researchers can carefully select excitation and emission filters specific to each target organism. Neu et al. [44] for instance developed a method allowing a clear distinction between signals from microalgae and cyanobacteria within photo-autotrophic biofilm. They presented a non-destructive multi-channel CLSM and 2P-LSM methodology allowing for differentiation of autofluorescent signals of cyanobacteria recorded in the red and far-red channels, as well as green algae recorded in the far-red channel only. However, they also indicate that this approach may be limited by confusing signal interpretation due to colour variations and by potential differences in the degree of overlayed signal caused by technical factors or cell characteristics (variation in amount or location of photosynthetic pigments).
Chlorophyll autofluorescence can also be employed for viability assessment and environmental toxicity. Nonetheless, non-viable cells might still exhibit fluorescence, making this characteristic alone an unreliable indicator for assessing viability. Hence, some authors have opted for the use of fluorescent stains to assess microalgae viability. It is imperative, however, to carefully select stains that do not overlap with the red signal emitted by chlorophyll [45]. One noteworthy example is the use of Propidium Iodide (PI) from the LIVE/DEAD *BacLight* viability assay kit [46], commonly employed for the characterisation of non-viable bacteria, which, due to its red fluorescence, presents challenges in accurately assessing non-viable microalgal cells [47]. As a result, researchers have turned to alternative green stains such as SYTOX Green (it stains DNA from membrane permeabilized cells), enabling the visualization of non-viable photosynthetic cells in green, thereby circumventing interference with chlorophyll autofluorescence [47-49].

As depicted in **Figure 1**, alongside the prevalent red autofluorescence of chlorophyll, several studies have reported the occurrence of distinct autofluorescence phenomena, including green fluorescence [50]. Initially considered a valuable taxonomic feature for characterizing dinoflagellates, green autofluorescence is now recognized as widespread among various photosynthetic organisms, encompassing dinoflagellates, diatoms, green algae, cyanobacteria, and raphidophytes. This green autofluorescence, observed in the cytoplasm of microalgal cells, exhibits heightened intensity following cell death or fixation procedures, a phenomenon that may be linked to pH shifts and exposure to substances like copper ions [50, 51]. While the occurrence, mechanism, and intrinsic compounds of emission of green autofluorescence are not yet fully characterized, some authors have suggested the involvement of degradation products of chlorophylls, while others suggest the role of flavonoids [50, 52, 53]. Notably, a prominent concern arising from the presence of this green autofluorescence is the potential interference signal it may cause to green-fluorescent stains.

## 4.2. Staining of non-photosynthetic organisms and EPS matrix

In assessing photosynthetic organisms, autofluorescence of chlorophyll serves as a valuable tool, however, for non-photosynthetic organisms like bacteria, staining with chemical fluorophores, genetically encoded fluorescent proteins or fluorogenic reporting systems becomes essential. Details of fluorescent dyes can be found elsewhere and will not be reiterated here [23, 54]. Instead, the focus will be on the overarching strategy and existing literature in application with photosynthetic biofilms. Labeling bacteria within photosynthetic biofilms can be achieved by introducing certain fluorochromes that bind to specific cellular components. For instance, in the study of Le Norcy et al. [55] investigating the anti-biofilm effect of Dibromohemibastadin-1, Syto9 a nucleic acid-binding dye, was employed to stain bacteria, enabling the simultaneous visualization of microalgae in red and bacteria in green within a mixed-culture. However, the use of such stains binding to nucleic acids alters the viability of organisms, necessitating the discarding of samples after image acquisition, and is
not adapted for real-time monitoring. Besides, this dye may not only stain bacterial DNA but also
nuclear and mitochondrial DNA in microalgae, as well as the nucleic acid-rich regions in
cyanobacteria. This overlap may complicate distinguishing signals in mixed biofilm systems. To
mitigate this, complementary strategies like counterstaining or genetic markers could improve
specificity. Spectral unmixing in confocal microscopy and 2P-LSM is another potential solution to
separate overlapping signals [56]. In non-axenic cultures, such fluorochromes will stain all bacteria
present in the sample, potentially limiting the ability to specifically pinpoint bacteria of interest - a
limitation that can be overcome by employing Fluorescence In Situ Hybridization (FISH) technology
[57]. As recently reviewed by Barbosa et al. [57], in combination with CLSM, the design of
fluorescently labelled, sequence-specific DNA probes, associated with *in situ* hybridization, enables
the spatial detection of specific microorganisms without disrupting the 3D structure of the biofilm.
Another alternative for specifically investigating the localization of specific bacteria is the use of
genetically encoded fluorescent proteins. In a study by Astafyeva et al. [58], researchers modified the
strain *Dyadobacter* sp. HH091, by implementing a plasmid carrying a GFP-encoded gene (green
fluorescent protein) along with a gentamycin resistance for plasmid selection, to assess its behavior
and role within a microalgal biofilm. As an example, **Figure 2** shows some CLSM images of
*Tetraselmis suecica* microalgae co-cultivated with the GFP-tagged bacteria, *Bacillus velezensis*
QST713-*gfp* in biofilm of 7 days. However, it is worth noting that the expression of these proteins
tends to diminish over time if no selection pressure is maintained to retain the plasmid, limiting the
potential for long-term kinetic experiments. Furthermore, as demonstrated in bacterial biofilms, the
use of a GFP reporter can be challenging due to specific requirements, such as the need for oxygen for
fluorophore development and the limitation of fluorescence at low pH [59, 60]. Currently, no study is
exploring the potential limitations that may hinder the application of GFP in microalgal biofilms,
which could be an interesting avenue for future research. Another crucial consideration when working
with photosynthetic biofilms is selecting staining methods that do not overlap with the chlorophyll
signal, as previously stated. While some researchers opt for alternative stains with distinct emission
signals, others prefer the use of autofluorescence quenching kits containing compounds that bind to
chlorophyll, thereby reducing fluorescence signals. For instance, the research conducted by Astafyeva
et al. [58], employed an autofluorescence quenching kit (TrueVIEW, Vector Labs, SP-8400) to
diminish the chlorophyll signal, enabling the observation of eGFP-tagged bacteria (enhanced green
fluorescent protein) in conjunction with microalgae. The authors mentioned that this kit contain a
hydrophilic compound able to bound electrostatically to chlorophyll, reducing its fluorescence. This
kind of quencher has been mainly used in immunofluorescent tissue analysis to lower signal-to-noise
ratios, thus enhancing the clarity of the target antigen signal [61]. This can represent a promising
strategy to overcome chlorophyll fluorescence to image photosynthetic biofilms and merits to be
tested.
EPS matrix plays pivotal roles as the structural foundation and communication hub [16]. To delve into the matrix composition, researchers often turn to fluorescent labelling with lectins [41, 62, 63]. Lectins, proteins with an affinity for specific carbohydrates, can be conjugated with fluorophores and used to visualize and study the intricate matrix architecture. A wide range of types of lectin-binding probes exists, each varying in their specificity for distinct carbohydrates. Importantly, organisms within biofilms might secrete different carbohydrates based on various factors, including individual biofilm constituents, environmental factors, or the age of the biofilms. Therefore, the effectiveness of lectin-binding may vary, emphasizing the need for careful consideration when selecting lectins, suitable for the experimental requirements. For example, Barranguet et al. [41] hypothesized that incomplete lectin-based staining of the EPS matrix in their study might be attributed to changes in EPS composition over time. As such, it is recommended to test specific microbial communities against a panel of lectins with different specificities to identify the most suitable one for accurate matrix composition analysis [64]. Similarly, as outlined in their study, the thickness of a biofilm can impede the diffusion of dyes used for matrix or bacterial staining, potentially compromising the effectiveness and precision of the methods. In another work, to surpass these challenges, a cocktail of lectins was used to stain the EPS matrix of biofilms of several photosynthetic organisms [39].

# 5. Confocal Laser Scanning Microscopy (CLSM)

## 5.1. Basic principles

CLSM is an advanced imaging technique that provides high-resolution, three-dimensional visualization of biofilms [65, 66], (Fig. 3, A). It involves the use of a focused laser beam to illuminate a specific plane within the biofilm sample. Unlike traditional microscopes, confocal microscopy selectively captures light emitted only from the focal plane, eliminating out-of-focus light. This is achieved through a pinhole aperture that allows only in-focus light to reach the detector, resulting in improved contrast and resolution. By scanning through multiple focal planes, a series of images can be reconstructed to generate a three-dimensional (3D) representation of the sample.

## 5.2. Advantages and limitations

CLSM stands as a cornerstone in visualizing biofilms, providing three-dimensional (3D) *in situ* images of living and fully hydrated microalgal biofilms [65]. This technique excels in differentiating individual cells and extracellular matrix components within the biofilm, shedding light on spatial distributions and chemical composition [67, 68]. Additionally, CLSM ability to perform real-time provides valuable insights into dynamic processes occurring within biofilms [69], as observed in **Figure 2**.

CLSM imaging involves a repetitive acquisition procedure and scanning through all spatial depths to compile a 3D image, resulting in volumetric imaging with extended acquisition times. Moreover, the
excitation light not only illuminates the specific volume of interest but also extends into a region above and below it, leading to constant illumination of a large volume of the sample all along the acquisition. This system may induce photobleaching and photodamage while unfortunately, this out-of-focus excitation is wasted and essentially discarded as the pinhole blocks these out-of-focus photons. Importantly, within a biofilm, photosynthetic organisms may exhibit a photokinesis or phototaxis response, involving their movement in response to varying light gradients [70]. While this phenomenon has not been extensively documented in the literature to our knowledge, it is crucial to highlight that the heightened light exposure from CLSM could potentially disrupt the organization of biofilm structures. Therefore, experiments involving CLSM should be executed swiftly to minimize potential alterations in the biofilm natural state. Two-photon laser scanning microscopy (2P-LSM) can also overcome some of these limitations by employing a powerful, pulsed infrared laser beam to enhance depth penetration during fluorescence excitation, with fluorescence generated within the focal volume [65]. This eliminates the need for a pinhole to reject out-of-focus fluorescence light, reducing photo-related phenomena compared to CLSM, although it can lead to a slight decrease in signal-to-noise ratio at similar imaging speeds due to slower signal generation rates and can also over-heat the sample. Caution related to loss of signal intensity should also be exercised when employing CLSM imaging, as several studies have reported its susceptibility to yielding unreliable data due to light attenuation [41, 71-74]. Indeed, as light penetrates deeper into the biofilm structure, it gets absorbed, scattered, or refracted, leading to reduced signal intensity and diminished clarity in imaging. In the study of Barranguet et al. [41], as the biofilm biomass increased, the transmitted light reaching the base of the biofilms decreased, with up to 90% of the incident light being absorbed by the biofilm. In another study, Fanesi et al. [74] demonstrated that light attenuation was linearly correlated with their mean thickness. As a result, the observation of thick biofilms may thus pose a technical challenge when using CLSM imaging, due to limitations in imaging depth and resolution, which hinder the accurate assessment of volumetric features [75]. In addition, the use of some supports brings additional challenges to the application of CLSM. For instance, fabric substrates, with their intricate fiber network, can obstruct light penetration and create shadowing effects, limiting imaging depth. The potential coloration of fabrics by certain stains is another factor to consider. Accordingly, **Figure 4** shows CLSM images of the microalgal biofilm *Tetraselmis suecica* grown on fabric substrates, highlighting co-staining of certain fibers and light reflection. Overcoming these substrate-specific constraints requires careful consideration of illumination and detection strategies. One possibility is the use of spectral analysis that helps to take in account precise excitation and emission spectra of fluorophores.

## 5.3. Current applications

CLSM is the most commonly used technique in biofilm research, and the field of photosynthetic biofilms is no exception. Most of this research focuses on deciphering the influence of biotic and
abiotic factors on biofilm structure. For instance, the species diversity plays a critical role in shaping biofilm formation and architecture, influencing structural properties and overall performance [9, 73, 76]. Irving and Allen [77] suggested that the development of microalgal biofilms is more significantly influenced by species selection and control than by support surface properties, as evidenced by the differing CLSM images of *Scenedesmus obliquus* and *Chlorella vulgaris*. In the study of Fanesi et al. [73], CLSM imaging observations showed that the architecture of microalgae biofilms is species-dependent, yet there are common developmental trends resembling those observed in bacteria and fungi. Initially, biofilms exhibit irregular surfaces with high roughness and thin structures, with some species having higher EPS volume compared to cell volume. In another study, Wang et al. [76] compared the relationship between biomass yield (mass divided by biofilm area : g/m²) and microstructures of co-cultured microalgal biofilms using CLSM (*Scenedesmus obliquus* co-cultured with *Chlorella pyrenoidosa*, *Scenedesmus acuminatua*, or *Ankistrodesmus falcatus*). They found that biomass yield decreases with lower biofilm porosity, and biofilms with smaller cell clusters and uniform microstructures exhibit significantly higher yields. They suggested that microalgal cells with smaller size, spherical shapes, and less EPS inhibit cell-cluster formation, thereby enhancing biomass production. Presence of bacteria may also play a critical role in microalgal biofilm formation [67, 68]. Through multimodal imaging - including CLSM, scanning electron microscopy (SEM), and time-of-flight secondary ion mass spectrometry (ToF-SIMS) - Yang et al. [67] found that bacterial quorum-sensing signal molecules (N-acyl homoserine lactones, AHLs) enhance diatom biofilm formation by increasing biomass, EPS content, and biofilm thickness, with effects influenced by carbon chain length and concentrations. Another study found that bacterial EPS production is significantly enhanced in the presence of algal exudates [68]. The bacterial transition from motile to sessile behavior induced by these exudates facilitates the formation of a joint extracellular matrix scaffold, thereby enhancing algal-bacterial aggregation. In addition to microbial species, shear stress has been proved to affect the mechanical properties of microalgal biofilms, as already reported for bacterial communities [74]. According to Fanesi et al. [74], high shear stress significantly increased biofilm cohesiveness, likely due to a biological response against hydrodynamics stress. Achieving an optimal balance between flow, with the resulting shear stress, and nutrient availability is crucial for high biomass production. Dense biofilms, often formed under minimal flow rate and shear stress, may face nutrient limitations, while elevated flow rates can enhance nutrient transport and growth, but may also increase cell detachment, impacting process stability. Interestingly, [78, 79] also showed that the light regime has a significant impact on the cell spatial organization of *C. vulgaris* biofilms. Indeed, while cells grow and spread evenly on the surface for low light, a clustering behavior was observed for higher light intensity, suggesting structural adaptation to light stress. Similarly, Morgado et al. [37] demonstrated that nutrient availability and light intensity had a significant impact on *Haematococcus pluvialis* biofilm growth in an rotating microalgal biofilm system. Another study had also shown that nitrogen deficiency induces irregular morphologies in photogranules, with the development of voids
and channels compared to nitrogen-replete conditions [10]. Despite nitrogen depletion, biomass productivity and performance were unaffected, a phenomenon likely linked to the increased surface area, which maximizes substrate contact and supply.

CLSM is a versatile tool that can be coupled with different biofilm-based cultivation system at pilot or laboratory scale. For instance at laboratory scale, Le Norcy et al. [80] combined CLSM with both microtiter plates and a microfluidic system to study the effects of anti-fouling compounds on the adhesion and growth of microalgal biofilms. This approach demonstrated that the two systems complement each other: microtiter plates are useful for initial static high-throughput screening, while microfluidic systems provide a more realistic simulation of natural conditions for evaluating anti-fouling effects. However, microtiter plates can suffer from issues such as evaporation and uneven light exposure, which can be addressed by filling peripheral wells with water, employing photon density cartography, and using black-walled, transparent-bottomed plates to improve light control. Another lab-scale cultivation method gaining traction in microalgal biofilm research, and compatible with CLSM, is 3D bioprinting, which enables the automated fabrication of 3D biological structures by integrating living cells with biocompatible polymers [81, 82]. For instance, Martin et al. [32] investigated the interaction between the green microalga, *Marinichlorella kaistiae*, and several bacteria including *Erythrobacter* sp., in an hydrogel system. They demonstrated an increase in growth rates within algal-bacterial hydrogels compared to hydrogels containing algae alone. Similarly, other authors demonstrated the potential for microalgae to grow with filamentous white-rot fungi in 3D-printed nanocellulose hydrogels [83]. Nevertheless, a major limitation of those systems is the light attenuation due to the self-shading of microalgae and biomaterial, limiting photosynthetic growth. That is why, Wangpraseurt et al. [84] developed a 3D printing management system inspired by coral morphology, mimicking features of living coral tissue and taking advantage of the high photosynthetic efficiency of the coral-microalgal symbiosis. This system incorporates structures that optimize light utilization supporting microalgal growth, showing promising applications in coral reef research and photobioreactor design. As such, by incorporating photosynthetic organisms into these printed structures, researchers can simulate and study complex interactions in a controlled laboratory setting. As this technology continues to evolve, it holds great promise for revolutionizing our approach to studying and harnessing the capabilities of photosynthetic biofilms.

On the whole, thanks to CLSM imaging coupled to different cultivating systems, a better understanding of the impact of operational factors (light, nutrients, hydrodynamics, microalgae strains) on biofilm properties and structural dynamics was achieved, helping ultimately biofilm-based process optimization.

# 6. Light-sheet microscopy

## 6.1. Basic principles
Light-sheet microscopes can capture a whole optical slice all in one go thanks to CCD (Charge Coupled Device) or CMOS (Complementary Metal Oxide Semi-Conductor) cameras. A single optical section is illuminated orthogonal to the detection axis [85, 86] (Fig. 3.C). It employs a thin plane of light that illuminates the sample only at the focal plane, thus reducing phototoxicity and photobleaching. This technique involves the intersection of two perpendicular objectives - one for illumination and one for detection - allowing for rapid three-dimensional imaging of large, living specimens with minimal photodamage.

## 6.2. Advantages and limitations

When compared to CLSM, this system facilitates the generation of 3D images from a large volume of the sample, bypassing the slow process of beam scanning through different depths, making the whole method much faster [87]. Moreover, illuminating a thin section of the sample, enables the excitation of molecules exclusively within the imaging plane. This means that only the target volume is illuminated, effectively reducing the risk of photodamage of the biofilm. As such, this technique is particularly valuable for capturing large-scale, 3D structures of biofilms, providing essential context for understanding their overall spatial organization and dynamic processes over prolonged durations. However, in contrast to CLSM, this improvement in speed and photon efficiency comes at the cost of slightly reduced image resolution. Besides, the complex three-dimensional structures of biofilms may not always align optimally with the imaging plane of the light sheet, potentially causing incomplete or distorted reconstructions [86]. Sample mounting in an orthogonal light sheet, albeit delicate, allows the rotation of the sample, so it opens to the possibility of orienting the sample and multi-angle acquisition of 3D samples.

## 6.3. Current applications

Light-sheet microscopy is widely applied for non-invasive, high-speed imaging in various biological research domains, including developmental biology, neuroscience, and cell biology [85, 86]. Despite offering clear advantages over CLSM, this technology has seen limited use in the study of bacterial biofilm structures, and to our knowledge, no studies have specifically focused on photosynthetic biofilms. A work by Qin et al. [88] successfully mapped individual cells of Vibrio cholerae trajectories as the 3D biofilm expanded, employing light-sheet microscopy in combination with cell-labeling to unveil the spatio-temporal patterns of development. In a recent study by Zhang et al. [89], a novel flow channel design was introduced, effectively addressing technical hurdles in studying biofilm growth via light-sheet technology, notably the need for the sample to be near the objective lenses. This advancement enables high-resolution imaging of thriving bacterial biofilms over prolonged durations, specifically utilizing lattice light-sheet technology. As such, given the continuous technological advancements and the benefits it offers including a decrease in photo-related phenomena, we could
anticipate a significant increase within photosynthetic biofilm research in the utilization of light-sheet technology in the upcoming years.

# 7. Histology coupled with a digital slide scanner

## 7.1. Basic principles

Histology methods, when coupled with a digital slide scanner, leverage the principles of tissue sectioning, staining, and digitization to provide detailed, high-resolution images of biological specimens [90] (Fig. 3.B.). This microscopic study involves a series of essential steps, beginning with the fixation of the tissue, to preserve its structure and prevent decay, by halting biochemical processes. Then, after tissue processing, embedding the specimen in a solid medium provides structural support, allowing the creation of sections that can be subsequently observed by a digital slide scanner, with the option of staining for analysis. This approach streamlines the generation of virtual slides, readily stored for convenient access, enabling researchers to navigate and analyse tissues at the cellular level with ease.

## 7.2. Advantages and limitations

Histology coupled with digital scanners offers benefits such as enhanced sharing and consultation of histologic images, improved storage, and preservation of stained slides [90]. This method represents a powerful tool for unraveling the complexities of biofilm structures, particularly within dense and thick biofilms on intricate substrates like fabrics with a complex network of fibers. By sectioning and examining a vertical 2D cross-section of the biofilm, it becomes possible to analyze cell distribution and architectural intricacies, with the possibility of quantitative data extraction. Furthermore, the scanner capacity for seamless image stitching facilitates the generation of precise, panoramic views of extensive biofilm areas, allowing the capture of high-resolution images of entire sample sections. This technique requires sample preparation and fixation, and while clearing of the tissue is often employed for immunolabeling, to our knowledge, such method has never been tested on biofilm. Sample clearing allows to explore in depth the 3D organization of fixed structures, previously opaque to photons. Many approaches have been described; among them it would be interesting to explore two main strategies: the use of iDISCO solvent that is very efficient on many tissue types. The use of iDISCO stiffens the sample that might be an advantage for mounting, but the sample size decreases limiting even more optical resolution of fluorescent labels. On the other hand, CUBIC aqueous solution is faster and maintains the gel-like aspect of the sample, but using some fluorescent dyes might be trickier. A recent attempt of biofilm clearing was published using an iohexol based solution (iCBiofilm) [91]. However, the requirement for sample fixation impedes the possibility of live imaging, limiting the dynamic observation of biofilm processes. Despite these drawbacks, the integration of histology with advanced imaging technologies remains a valuable asset in biofilm
research, shedding light on intricate structural details and contributing to our understanding of biofilm
behavior in diverse contexts.

**7.3. Current applications**

This technology finds extensive application in imaging whole-slide tissue sections, primarily in
pathology and histology studies [90]. It has also been employed in biofilm research, notably for
monitoring bacterial pathogen biofilm development in live organism tissues [92, 93]. Nonetheless, to
our knowledge, this technique has not yet been described to image photosynthetic biofilms. However,
as we demonstrate here (Fig. 3.B), its potential value arises when investigating biofilms grown on
complex porous surfaces, as it enables the detailed observation of cell attachment and biofilm
development across the entire structure.

**8. Optic Coherence Tomography (OCT)**

**8.1. Basic principles**

Optical Coherence Tomography (OCT) operates on the principles of low-coherence interferometry, utilizing near-infrared light to generate high-resolution, cross-sectional images of biological tissues or materials [94] (Fig. 3.D.). By measuring the echo time delay of backscattered light, OCT creates detailed, non-invasive images with micron-scale resolution, enabling depth profiling and real-time visualization of structures.

**8.2. Advantages and limitations**

Compared to CLSM, OCT can be used to characterize large sample volumes and biofilm mesoscale
structures with high resolution [95]. Compared to microscale structure covering an area of 10-100 µm
biofilm surface, the mesoscale covers several mm large [95]. By applying, low-coherence light waves
onto the sample surface, OCT captures the backscattered light to create high-resolution cross-sectional
biofilm images, providing non-invasive visualization of biofilm layers and their structural variations
over time. Its non-destructive nature also enables longitudinal studies, offering a unique perspective on
biofilm development and response to environmental changes. However, although OCT excels in
providing structural details, its limitation lies in the inability to distinguish between biofilm
constituents, which sets it apart from other imaging techniques that may offer more comprehensive
functional information.

**8.3. Current applications**

This technology has found widespread applications in medical imaging, or ophthalmology, providing
valuable insights into tissue morphology and aiding in the diagnosis and monitoring of various
conditions. Today, OCT has found a niche in the study of biofilms due to its non-invasive, label-free
approach [75, 95, 96]. Several studies have successfully used OCT imaging to monitor biofilm
formation over time, allowing temporally resolved profiling and spatial resolution with observation of
cavities and porosity [97-99]. In particular, Fabbri et al. [100] introduced a novel approach utilizing real-time mesoscale OCT imaging for the assessment of marine biofilm fouling on biocidal antifouling coatings. They developed a novel marine fouling flow cell with a clear plastic lid, allowing for non-destructive, real-time OCT imaging. The authors suggested that cross-sectional imaging enables detailed analysis of biofilm thickness which might be more predictive of drag forces than surface coverage. However, the authors also noted that the approach is time-consuming and not suitable for high-throughput testing. Additionally, due to the small size of the flow cell, it is rather suitable to test biofouling by small microorganisms than larger ones. While recent studies demonstrated the interest of applying OCT imaging to characterize photosynthetic biofilms, very few concerns ecological applications. Among them, Depetris et al. [101] combined OCT imaging with automated oxygen microprofiling and amplicon sequencing to respond to key questions regarding the structure-function of phototrophic biofilms in streams and rivers. Their work highlighted the role of hydrodynamics, that allows for advective transport around and within biofilms patches, and that induces metabolic shifts at the scale of the overall biofilm. Lamprecht et al. [102] also pointed out the contribution of the periphyton establishment to changes in structural properties of synthetic freshwater communities.

From the aforementioned, we believe that the extensive use of OCT imaging to monitor, lab-scale to large scale, photosynthetic biofilm cultures would bring a deeper insight into the relationship between process productivity and stability and biofilm architecture. Although OCT proves to be a valuable tool, it lacks the capacity to characterize biofilm constituents, a task better suited for CLSM. Hence, a combination of both methods is complementary, allowing for a more comprehensive assessment of biofilm structures at different scales. For the first time, Fanesi et al. [39] used the combination of CLSM and OCT to successfully characterize the architectural features of monospecific microalgal biofilms, bridging the gap between the micro- and mesoscale. In particular, they introduced a customized flow-through system utilizing a spectrophotometer cuvette as a chamber. This configuration enables a thorough analysis and screening of biofilm traits through CLSM and OCT, ranging from metabolic activity to microscopic structures. As such, this versatile tool encompasses not only structural examination but also real-time physiological assessments, providing additional insights and enhancing our understanding of microalgal biofilm behavior.

## 9. Conclusion

Real-time and non-destructive investigation of microalgal biofilm development has become possible thanks to advances in photonic imaging techniques. Coupled with lab-scale cultivation using flow-through systems, microtiter plates and 3D bioprinting, it is now possible to decipher biofilm structural dynamics in non-static and static conditions, as well as inherent microbial interactions, to optimize biofilm-based technologies. Such a study involves a strategic combination of autofluorescence and staining techniques. Autofluorescence, driven by the inherent pigment contents, such as chlorophyll, provides a valuable tool for non-destructive imaging and differentiation of photosynthetic organisms.
Complementary staining methods, including the use of chemical fluorophores or genetically encoded fluorescent proteins, enhance our understanding of spatial distribution of non-photosynthetic organisms and the EPS matrix. However, researchers must remain vigilant about potential limitations, such as interference between stains and autofluorescence signals, as well as limited diffusion in thick biofilms, to ensure the accuracy and reliability of their methods. The selected imaging techniques in this review, CLSM, light-sheet microscopy, and histology coupled with digital slide scanners, and OCT, each offer specific advantages for dissecting the intricate dynamics of microalgal biofilms. These approaches face a fundamental trade-off between high speed and 3D imaging capability, often sacrificing one for the other, along with compromises in signal-to-noise ratio, photo-related phenomena, structural information or real-time analysis. However, taken together, they are complementary tools, providing advancements in our understanding of these dynamic ecosystems and paving the way for innovative applications in biotechnology and environmental engineering.

## Acknowledgments

This work was funded by the French ANR "PhotoBiofilmExplorer" and the Université-Paris-Saclay Biosphera program. We would like to thank Andrea Fanesi from CentraleSupélec LGPM for his precious help all along the project, as well as Pierre Adenot from the INRAE MIMA2 imaging facility for acquiring the illustrative light-sheet images (doi.org/10.15454/1.5572348210007727E12) and Julie Rivière from the INRAE Abridge platform for histologic analysis.

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Figures

<figure><img src="image_3.png" /></figure>

**Figure 1:** CLSM observations of microalgal biofilms: left *Cylindrotheca closterium* AC170; right, *Tetraselmis suecica* AC254. Microalgae exhibiting red and green autofluorescence can be observed. White scale bars represent 20 µm. Arrows point out cells with green autofluorescence as examples.

<figure><img src="image_4.png" /></figure>

**Figure 2:** 3D reconstruction of co-cultured microalgal-bacterial biofilms obtained from CLSM z-stacks using IMARIS software. In green, *Bacillus velezensis* QST713-gfp expressing GFP is depicted, while in red, *Tetraselmis suecica* AC254 is highlighted. Biofilms were cultivated in 96-well microtiter plates for 7 days at room temperature under homogenous and continuous LED light. Three microalgal concentrations (T1 = 10⁴ cells.mL⁻¹; T2 = 10⁵ cells.mL⁻¹; T3 = 10⁶ cells.mL⁻¹) and three bacterial concentrations (B1 = OD₆₀₀nm 0.05; B2 = OD₆₀₀nm 0.1; B3 = OD₆₀₀nm 0.5) were mixed in
microalgal culture media prepared with synthetic sea salt Instant Ocean (Aquarium Systems, Mentor, OH); 80 µL of medium were removed and replaced with fresh medium each day. CLSM acquisition was performed using a Leica HCS-SP8 Microscope (CLSM, LEICA Microsystems, Wetzlar, Germany) with a x40 objective. White scale bars represent 30 µm. Image 3D reconstruction was performed using IMARIS software (ver. 9.3.1, Bitplane AG, Zurich, Switzerland).

<figure><img src="image_5.png" /></figure>
<figure><img src="image_6.png" /><figcaption>**Figure 3: Photonic imaging of microalgal *Tetraselmis suecica* AC254 biofilm cultivated on fabric support.** Schematic details of microscopy observation strategies are provided, (adapted with authorizations from Guéneau et al. [103]). **A. Confocal Laser Scanning Microscopy (CLSM) imaging.** On the left, there is a mosaic view with zoomed images of biofilm, and on the right a</figcaption></figure>
representative IMARIS easy3D projections of z-stack images using IMARIS software (ver. 9.3.1, Bitplane AG, Zurich, Switzerland). The shadows on the left represent the virtual lateral biofilm projection of the submerged biofilm. **B. Histology coupled with digital slide scanner.** Here is a 2D vertical section of the biofilm with the support. After fixation and staining, entire sections were digitalized using a Hamamatsu NanoZoomer (Tokyo, Japan). For **A.** and **B.** microalgae can be visualized in red, and in green are the bacteria along with the fiber fabric, colored with Syto9. **C. Light-sheet microscopy.** On the left, various 3D views of a biofilm sample are presented, with the biofilm fluorescent signal depicted in red. On the right, a zoomed 3D representative image of a top-view is shown, obtained from z-stacks. The color scale represents various depths; red indicates the top cells within the biofilm structure, and dark blue represents the bottom cells. **D. Optical coherence tomography (OCT)** image showing a vertical 2D cross-section of a microalgal biofilm.

<figure><img src="image_7.png" /></figure>

**Figure 4: CLSM imaging of microalgal biofilm *Tetraselmis suecica* AC254 grown on fabric substrates.** **A.** Observation of photosynthetic biofilms at different scales. **B.** Representative IMARIS easy3D projections of CLSM z-stack images using IMARIS software (ver. 9.3.1, Bitplane AG, Zurich, Switzerland). The shadows on the right represent the virtual lateral biofilm projection of the submerged biofilm. Microalgae are depicted in red, blue corresponds to the laser reflection from the fibers, and green denotes the bacteria and staining of fibers with Syto9. As observed, the fabric substrate is composed of two distinct fibers with different chemical and physical properties.
<table><caption>Table 1: Comparison of the different imaging techniques for application in microalgae biofilm observations.</caption><thead><tr><td>Technique</td><td>Resolution</td><td>Advantages</td><td>Disadvantages</td><td>Applications</td></tr></thead><tbody><tr><td>Confocal Laser Scanning Microscopy (CLSM)</td><td>High (submicron)</td><td>High 3D resolution, ability to differentiate biofilm components (cells, polymers), real-time, compatible with multiple culture systems</td><td>Light attenuation, limited depth photobleaching, slow acquisition</td><td>Visualizing spatial organization, components distribution, biofilm architecture, studying microbial interactions in biofilms</td></tr><tr><td>Light-sheet Microscopy</td><td>Moderate to High (micron scale)</td><td>Rapid 3D volumetric imaging, reduced photodamage, large-scale biofilm imaging</td><td>Limited resolution, incomplete or distorted reconstructions of complex structures. Complexity of mounting: embedding in a gel, mounting in a tube, orientation of the sample during the scan, access to the sample with the objective.</td><td>Mapping large-scale biofilm structure</td></tr><tr><td>Histology + Digital Slide Scanner</td><td>High (2D sections)</td><td>Detailed spatial organization, excellent for thick biofilms on complex substrates</td><td>Requires fixation, not suitable for live imaging</td><td>Structural analysis of dense biofilms, studying biofilm growth on porous materials</td></tr><tr><td>Optical Coherence Tomography (OCT)</td><td>Moderate (> micron scale)</td><td>Non-invasive (no dyes), real-time imaging, mesoscale structural analysis</td><td>Cannot distinguish biofilm components, limited functional information</td><td>Profiling biofilm thickness, studying mesoscale structure</td></tr></tbody></table>
**Credit authors statement**

JM wrote the original draft; JM, AS, JD and VC acquired the microscopic images; JM, RB and FL reviewed & edited the manuscript; RB and FL supervised the project and acquired the financial support for the project.

**Declaration of interests**

[x] The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

[ ] The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:
Highlights

* Techniques like CLSM, light-sheet microscopy, histology coupled with a digital slide scanner, and OCT provide unprecedented insights into the structure and function of microalgal biofilms.

* Photonic imaging enables real-time, non-destructive monitoring of biofilm development, crucial for optimizing biofilm-based cultivation.

* Combining autofluorescence with staining techniques improves the study of photosynthetic and non-photosynthetic components within biofilms.

* The field requires common protocols and benchmarks to ensure consistency and reliability in photosynthetic biofilm imaging studies.