Microalgal biofilms: A further step over current microalgal cultivation techniques

Science of the Total Environment 651 (2019) 3187–3201

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Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv

Review

Microalgal biofilms: A further step over current microalgal cultivation techniques Antonia Mantzorou, Filippos Ververidis ⁎ Plant Biochemistry and Biotechnology Group, Biological and Biotechnological Applications Laboratory, Department of Agriculture, School of Agriculture, Food and Nutrition, Technological Educational Institute of Crete, Heraklion, Greece

H I G H L I G H T S

G R A P H I C A L

A B S T R A C T

• Currently, microalgal cultivation in large scale has been limited. • Biofilms are different life forms where microorganisms grow attached to substrata. • Biofilms could resolve the bottlenecks of suspended cultivation systems.

a r t i c l e

i n f o

Article history: Received 6 July 2018 Received in revised form 24 September 2018 Accepted 28 September 2018 Available online 1 October 2018 Editor: Henner Hollert Keywords: Microalgae Suspended cultivation systems Biofilms

a b s t r a c t The scientific community has turned its interest to microalgae lately, because of their countless applications such as wastewater treatment and pharmaceutical industry. Nevertheless, so far applied cultivation methods are still prohibitive. Ordinary cultivation techniques in which microalgae are suspended in liquid medium suffer from many bottlenecks, such as low biomass productivities, difficulty in biomass harvesting and recovery, high installation and operating cost, high water requirements etc. Although, microalgal biofilms are known to be a nuisance because of surfaces fouling, they have emerged as an innovative technology with which microalgae are developed attached to a solid surface. This technique seems to be advantageous as compared to conventional cultivation systems. Microalgal biofilm systems could resolve the problematic aspects of ordinary cultivation techniques such as low biomass productivities, water management and biomass recovery. A detailed description of this technique with respect to the parameters affecting them is reviewed in this work. © 2018 Published by Elsevier B.V.

Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3188 Microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3188

⁎ Corresponding author at: Technological Educational Institute of Crete, P. O. Box 1939, GR-710 04 Heraklion, Crete, Greece. E-mail address: [email protected] (F. Ververidis).

https://doi.org/10.1016/j.scitotenv.2018.09.355 0048-9697/© 2018 Published by Elsevier B.V.

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3. 4. 5.

Conventional cultivation techniques and recovery of microalgal biomass Microalgal biofilms . . . . . . . . . . . . . . . . . . . . . . . . Main microalgal based biofilm cultivation systems . . . . . . . . . . 5.1. Permanently immersed biofilms . . . . . . . . . . . . . . . 5.2. Biofilms between two phases . . . . . . . . . . . . . . . . 5.3. Permeated biofilm systems . . . . . . . . . . . . . . . . . 6. Factors affecting microalgal biofilms . . . . . . . . . . . . . . . . 6.1. Selection of proper microalgal strain . . . . . . . . . . . . . 6.2. Nutrients . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. Light availability . . . . . . . . . . . . . . . . . . . . . . 6.4. Temperature . . . . . . . . . . . . . . . . . . . . . . . . 6.5. pH . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6. CO2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.7. Attaching material . . . . . . . . . . . . . . . . . . . . . 6.8. Flow velocity . . . . . . . . . . . . . . . . . . . . . . . . 6.9. Presence of other microorganisms . . . . . . . . . . . . . . 7. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction The considerable potential of microalgae for biomass production and co–products has been extensively examined lately from many scientific aspects of research (Berner et al., 2015). Although microalgae seem to be organisms with a range of quite different applications, there is a need in boosting their quantity and at the same time, keeping the cost at low levels (Mantzorou et al., 2018). Until now, the microalgal cultivation in large scale is not quite popular as it cannot become economically viable due to high installation and operating costs as well as low productivities (Berner et al., 2015; Gross et al., 2013). Most studies related to microalgal cultivation, have focused in suspended cultivation systems (Toninelli et al., 2016). Until now, there is a significant research aiming to improve the currently used cultivation systems by either improving process design or increasing biomass yield (Berner et al., 2015). Attached cultivation systems based on biofilms seem to be an attractive process because such systems have less water and energy requirements (Ozkan et al., 2012). Microalgal biofilms have been studied from both technological and ecological aspects (Zippel et al., 2007). Applications of these biofilms, such as aquaculture, wastewater treatment and improvement of antifouling substances, have gained the scientific interest (Roeselers et al., 2007). Nevertheless, as compared to bacteria involvement to fouling, knowledge about microalgae fouling is rather limited (Zippel et al., 2007). The purpose of this work is to revise the current literature concerning microalgal biofilms development as a new technology. This review attempts to present the current status of existing research data showing the important factors and synergistic effects of biofilm creation either oriented to marine biofilms or to freshwater systems. Taking into consideration of the latter's scarcity for human consumption, it is obvious that the cultivation of marine microalgal species is more encouraged than the freshwater microalgal cultivation (Khan et al., 2018). Experimental biofilm cultivation systems are described below with respect of their selected parameters, to give an insight of the contemporary trends in improving microalgae production. Factors affecting their performance are analyzed below for identifying the gaps of the ongoing research. 2. Microalgae Microalgae are normally unicellular microorganisms possessing a capacity to aggregate and thus being able to form different types of cell organization, such as unicellular, colonial and filamentous (Arad and Richmond, 2007; Palma et al., 2017). They are considered being

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the earliest life–forms in the earth (Hamed, 2016). They are thallophytes i.e. they are plants without leaves, roots or stems and chlorophyll a is the main photosynthetic pigment (Dragone et al., 2010). Their photosynthetic mechanism is similar to terrestrial plants but because of their simpler cell structures and their easiness of accessing to CO2 and nutrients, they have greater efficiency in converting solar energy into biomass (Priyadarshani et al., 2012). They live in a variety of environmental conditions and they are found in both aquatic and terrestrial environments. They grow under a broad range of temperature and their raise could be confined because of limited nutrient and light supply (Alam et al., 2015; Hannon et al., 2010). Those tiny microorganisms are of high importance due to their influence in the various existing ecosystems (Katarzyna et al., 2015; Sevda et al., 2017). As primary producers of the food chain, they are the most important level in the maintenance of trophic equilibrium (Monteiro et al., 2011). Microalgae have gained the scientific interest lately, since they have been characterized as a raw material for processing or manufacturing industry (Gross et al., 2013; Katarzyna et al., 2015). The applications of these microorganisms seem to be countless. Some of them are the fourth-generation biofuels, fertilizers, aquaculture feed, nutraceutical and wastewater treatment (Gross et al., 2013; Irving and Allen, 2011; Katarzyna et al., 2015; Zippel et al., 2007). Furthermore, because of their miscellaneous environmental habitats, microalgae can be used in ecotoxicological tests, for monitoring of water quality (Campanella et al., 2001; Garbowski et al., 2017). The cultivation of microalgae is preferable as compared with the terrestrial plants due to the reasons mentioned below. I) They have variable chemical compositions which are dependent on their cultivation medium. This is a result of ensuing from their huge biomass biodiversity (Choudhary et al., 2017). II) They do not compete with the terrestrial plants for agricultural land as they do not need any of it for their cultivation (Johnson and Wen, 2010). III) Their nutrients and water requirements can be met with the use of wastewater (Garbowski et al., 2017; Zhang et al., 2018). IV) Their cultivation has no seasonal limitations and some species survive in extreme environmental conditions. The fact that they double their biomass in a few hours results in an important rise in yields (Sevda et al., 2017; Wu et al., 2012). V) They cause less environmental pollution because of their less need for pesticides and fertilizers (Das, 2015). VI) Also, they could remove pollutants (such as nitrogen, phosphorus and heavy metals) from liquid waste streams and CO2 from the atmosphere, providing the extra benefit of phycoremediation (Hu et al., 2008; Palma et al., 2017; Wu et al., 2012). VII) Some microalgal species produce greater amount of bioenergy per area than the conventional oil crops do (Schenk et al., 2008; Schnurr et al., 2013; Shen et al., 2018).

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Fig. 1. Simplified designs of microalgal suspended cultivation systems: [a] raceway pond; [b] circular pond; [c] helical PBR; [d] vertical PBR; [e] flat–plate PBR.

3. Conventional cultivation techniques and recovery of microalgal biomass Currently, microalgae are cultivated in open systems or in closed systems (Photobioreactors–PBRs). These two system types could have various configurations (Fig. 1). In both cases, the microalgal cells are suspended in liquid culture media or water with the latter being the

basic component of the microalgal cultures (Berner et al., 2015; Cheng et al., 2013; Liu et al., 2013; Ozkan et al., 2012). It has been calculated that to produce 1Kg dry biomass of microalgae, about 12–2000 kg of water are needed (Berner et al., 2015). Both systems are compared in Table 1. A major problem in production of microalgae in large scale is the harvesting techniques. Because of their fast growth, their repeated

Table 1 Comparison of the most common microalgal cultivation techniques. Characteristics Open ponds

Advantages

Disadvantages

Closed systems

Advantages

Disadvantages

Reference Easy structure and operation Low capital and operation cost Easy, combined operation with wastewater treatment Free solar illumination Difficulty in handling contaminations Lower biomass productivity than closed systems Light limited system High water requirements due to low biomass/water ratio Cost of water transportation Water loss because of evaporation Difficulty in biomass harvesting Cost for biomass dewatering Dependent on environmental conditions Settling in big land area Insufficient mixing Control of environmental parameters pH, temperature, light intensity Appropriate for single microalgal species Minimized contaminations Better light availability Difficulty in designing bioreactor systems Low biomass productivity Difficulty in biomass harvesting High water requirements due to low biomass/water ratio Cost of water transportation High energy and operation cost Potential hydrodynamic stress of culture cells

Garbowski et al., 2017; Lee et al., 2014 Christenson and Sims, 2011; Lee et al., 2014; Sevda et al., 2017 Garbowski et al., 2017 Kumar et al., 2015 Garbowski et al., 2017; Katarzyna et al., 2015 Christenson and Sims, 2011; Lee et al., 2014 Hannon et al., 2010; Lee et al., 2014; Sevda et al., 2017 Berner et al., 2015; Sevda et al., 2017 Berner et al., 2015 Garbowski et al., 2017; Sevda et al., 2017 Berner et al., 2015; Lee et al., 2014 Berner et al., 2015 Dragone et al., 2010; Sevda et al., 2017 Sevda et al., 2017; Shukla et al., 2017 Shukla et al., 2017 Kumar et al., 2015; Sevda et al., 2017 Garbowski et al., 2017; Sevda et al., 2017 Hannon et al., 2010 Sevda et al., 2017 Sevda et al., 2017 Berner et al., 2015 Berner et al., 2015; Lee et al., 2014 Berner et al., 2015 Berner et al., 2015 Hannon et al., 2010; Sevda et al., 2017 Garbowski et al., 2017

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Table 2 Comparison of the most common microalgal harvesting techniques used in suspended cultivation systems. Characteristics Flocculation

References Advantages Drawbacks

Centrifugation

Advantages

Drawbacks

Filtration

Advantages

Drawbacks

Ultrasound

Advantages

Drawbacks Flotation

Advantages

Drawbacks

Electrophoresis

Advantages

Drawbacks

High efficiency of some flocculants e.g. chitosan Relatively fast method Precaution in appropriate selection of flocculants Chemicals required Sensitive to pH changes No chemicals required Rapid separation Good biomass recovery High energy consumption Expensive equipment Inappropriate for large culture volumes Possible cell rupture because of shear and gravitational forces Cost effective technique No chemicals required A variety of different membrane designs is available Slow process Membrane clogging and fouling Need for pre–concentration step Appropriate only for large cells It does not cause any shear stress to microalgal cells Non fouling method Low operation space At this time in progress Aggregation of other sediments than microalgal cells such as mercury No chemicals required Rapid method Low cost of initial equipment Low operation space Proven to be suitable only for small scale microalgae Uncertainty about its economical and technical sustainability High operational cost Cost effective No chemicals required Eco–friendly Cathode fouling and decrease of current intensity due to its reuse Change of cell composition due to high current densities

harvesting is recommended otherwise photosynthesis will be limited (Garbowski et al., 2017). Furthermore, due to the small size of microalgal cells (more than a few micrometers), recovery of the biomass becomes difficult and costly (Garbowski et al., 2017; Irving and Allen, 2011; Lee et al., 2014). Many authors have reported that harvesting techniques is the determinant factor for the cost assessment of microalgal cultivation (Garbowski et al., 2017; Irving and Allen, 2011; Lee et al., 2014; Miranda et al., 2017; Shah et al., 2014). Among them, the most popular harvesting techniques are centrifugation, flocculation, flotation, sedimentation and filtration (Garbowski et al., 2017). The cost of such methods accounts for the 30% of the total expense of microalgal cultivation (Irving and Allen, 2011; Lee et al., 2014). The proper harvesting technique, which one may choose, is contingent on microalgae features such as culture density, cell size and production purpose (Brennan and Owende, 2010). Nevertheless, none of them has been characterized economical and appropriate for microalgal production in large scale (Muñoz et al., 2009). These techniques are compared in Table 2. 4. Microalgal biofilms Biofilm could be characterized as consortium of microorganisms, embedded in extracellular polymeric substances (EPSs), forming a complex structure which is developed in solid surfaces (substrata) (Mantzorou et al., 2018). Biofilms are an alternative approach of microorganism communities which have totally different characteristics than those of suspended cultures (Berner et al., 2015). Despite the limited research, biofilms are not a new phenomenon. The earliest fossil biofilm that has been recorded, dates 3.5 billion years ago (Roeselers et al., 2008). Nevertheless, Costerton and his coworkers introduced the term biofilm in order to describe a consortium of organisms design, surrounded of extracellular matrix and attached to a

Suali and Sarbatly, 2012 Al hattab et al., 2015 Suali and Sarbatly, 2012 Brennan and Owende, 2010 Shukla et al., 2017 Suali and Sarbatly, 2012 Barros et al., 2015; Dragone et al., 2010 Suali and Sarbatly, 2012 Barros et al., 2015; Suali and Sarbatly, 2012 Suali and Sarbatly, 2012 Chen et al., 2011 Dragone et al., 2010 Suali and Sarbatly, 2012 Garbowski et al., 2017 Milledge and Heaven, 2013 Suali and Sarbatly, 2012 Barros et al., 2015; Ghosh and Das, 2015; Shukla et al., 2017 Dragone et al., 2010 Brennan and Owende, 2010 Bosma et al., 2003 Brennan and Owende, 2010 Bosma et al., 2003 Suali and Sarbatly, 2012 Suali and Sarbatly, 2012 Brennan and Owende, 2010 Barros et al., 2015 Ghosh and Das, 2015 Barros et al., 2015 Al hattab et al., 2015 Brennan and Owende, 2010 Al hattab et al., 2015; Shah et al., 2014 Al hattab et al., 2015; Chen et al., 2011 Chen et al., 2011; Ghosh and Das, 2015 Al hattab et al., 2015; Chen et al., 2011 Al hattab et al., 2015 Al hattab et al., 2015

substratum in contact with a liquid medium (Polizzi et al., 2017). In the aquatic environment, they cover most of the lower solid layers such as ships, rocks, marine animals etc., (Krishnan et al., 2017). They are usually composed with protozoa, bacteria, larvae and microalgae (Katarzyna et al., 2015). In some cases, it is possible to include higher organisms such as macroalgae (Berner et al., 2015). Biofilms, composed of multiple species are often used in wastewater treatment while biofilms composed only of single species are used in manufacturing process (such as biotransformations, fermentations etc.) (Li et al., 2007). Biofilms can be found in nearly every liquid–solid interface existing in nature and play a crucial role in ecosystems function (Li et al., 2007). They contribute considerably to nutrient movement or exchange but also to the energy flow (Zippel and Neu, 2005). Many scientific fields such as engineering, ecology, architecture and biology have investigated biofilms (Berner et al., 2015). Biofouling is the phenomenon by which all natural and artificial submersed surfaces are colonized by a consecution of organisms (Mieszkin et al., 2013). Until recently, biofilms have been characterized as a problem in fouling surfaces as they cause alloys and metal corrosion (Irving and Allen, 2011; Krishnan et al., 2017). In addition, they damage systems related to water distribution, power plants and cooling systems. In the last case, biofilms raise the pressure drop in heat exchangers and diminish the thermal efficiency (Sekar et al., 2004). Within several minutes, this complex phenomenon takes place and the solid surfaces are covered with organic and inorganic substances. Hence, the physico–chemical features of the surfaces are modified and subsequently the settlement of further microorganisms such as protozoa, bacteria etc., is influenced (Krishnan et al., 2017). Microalgal biofilms include all types of particular structures that consist of microalgae and bacteria which are the main organisms that compose them. These biofilms could be found in solid substrata adequately

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humidified, illuminated and capable to supply the microorganisms with nutrients. Many other organisms could be included in microalgal biofilms (non–axenic cultures) like bacteria which play a crucial role in the biofilm formation (Mantzorou et al., 2018). Microalgal biofilms are also identified as autotrophic biofilms consisting of microalgae (including cyanobacteria) and heterotrophic microorganisms (fungi, bacteria and protozoa) (Choudhary et al., 2017; Sukačová et al., 2015). The formation of microalgal biofilms is considered to be a complicated procedure and still not sufficiently understood (Katarzyna et al., 2015; Zeriouh et al., 2017). It is also believed that the process of biofilm formation and growth is different and depended on the species that take part in. Hence, it seems difficult to make a point about the potential process for the various types of existing biofilms (Choudhary et al., 2017). Nevertheless, the process of microalgal biofilm formation can be divided into two steps. In the first step, the cells initially adhere to the solid substratum through adsorption in order to form a conditioning film. This step is usually reversible. In the second step, a second irreversible adhesion follows because of the EPSs production (Shen et al., 2014b). Proteins, cations and organic molecules, concentrated on surfaces, favor the growth of microorganisms and the formation of such biofilms (Christenson and Sims, 2011). The biofilm further grows by cell division of microorganisms which are involved in the whole process instead of captivating free suspended particles from the nearby environment (Katarzyna et al., 2015). The biofilm gradually matures and its height raise creating a three dimensional structure with multiple layers. The cells are immobilized and cell clusters are formed. Microalgae are fed with nutrients which are moved via diffusion (Berner et al., 2015). Organic substances which are produced photosynthetically are available to the biofilm structure. These compounds will be degraded by bacterial communities (Zippel and Neu, 2005). The biofilm maturity influences the abundance and the proportion of participating organisms and EPSs (Schnurr and Allen, 2015). Afterwards, at a given biofilm thickness, losses (due to cell death, grazing, parasitism etc.) become almost equal to the growth. When the loss phase overdraws the growth phase, the growth of biofilm structure decreases (Boelee et al., 2014b). Microalgal cells on biofilm are able to acclimatize themselves in different environmental conditions (Kesaano and Sims, 2014). Besides, the biofilm formation is considered to be a protected growth style which provides to the microorganisms advantages concerning their survival in hostile environments (Li et al., 2007; Palma et al., 2017). It has been shown that bacteria in biofilm forms, resist 100–1000 fold to antimicrobial substances (Nithya et al., 2014). Guasch et al. (2016) examined the effect of triclosan and snail grazing to diatom community, growing in sandblasted glasses. They showed that in opposition to the grazer's presence, under triclosan, the diatom composition of the biofilm changed. Also, phosphorus uptake capacity of microalgae was diminished in the treatments with triclosan whereas the opposite was observed in the diatoms treated with grazers. In non–axenic conditions, various organisms can be concentrated in areas which are favorable for them. This leads to the creation of a heterogeneous biofilm structure. For example, cyanobacteria could be aggregated at the top of the structure, whereas anaerobic and heterotrophic bacteria settle in deeper layers, where oxygen and light are limited (Berner et al., 2015). The cells of various microorganisms (such as microalgae and bacteria) secrete EPSs which help them to be attached and stabilized on the surfaces (Choudhary et al., 2017; Zippel et al., 2007). The EPSs are composed with high weight molecules such as proteins, phospholipids, polysaccharides and nucleic acids and they include functional groups such as phosphoric, carboxylic, hydroxyl and amino groups (Christenson and Sims, 2011; Shen et al., 2015). EPSs are essential for the biofilm formation because they keep the biofilm together (Roeselers et al., 2008). Also, they help the microorganisms to stay attached to the solid surfaces and the particular matter such as erosion products, to be retained in the structure (Bott, 2011; Katarzyna et al., 2015). Furthermore, the formed EPS matrix could be considered as a storage space for water and nutrients and it has a protection role for

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the cells against extreme environmental conditions (e.g. pH and temperature fluctuation, dehydration) and harmful chemicals (Berner et al., 2015; Schnurr and Allen, 2015). EPSs are influenced by various factors such as nutrient availability, stress response, biofilm age and species composition (Choudhary et al., 2017). It is important to note that there are microalgal species (such as Chlorella vulgaris) incapable to produce EPSs by themselves (Katarzyna et al., 2015). It has been reported that the growth of Chlorella vulgaris alters from planktonic to biofilm when other species are present (Irving and Allen, 2011). Nevertheless, Gross et al. (2013) noticed that Chlorella vulgaris cells in axenic conditions, did managed to grow attached to a cotton sheet surface. In addition, it has been shown that the EPS production is enhanced by increasing light intensity and temperature (Kirsten and Lucas, 2002). During the last decades, the study of biofilm based cultivation of microalgae has gained considerable attention lately (Polizzi et al., 2017; Toninelli et al., 2016). This interest has recently emerged due to the need of a better and more economically harvesting and dewatering algae system (Christenson and Sims, 2011; Gross et al., 2013). Microalgal biofilms could be considered as an alternative system for biomass production. They could resolve the problematic aspects of suspended cultivation systems and meet the needs of biofuel production and wastewater treatment (Berner et al., 2015; Kesaano and Sims, 2014). Their advantages are mentioned below. 1. The water requirements are less than those of suspended cultures (Podola et al., 2017; Shukla et al., 2017). In suspended cultivation systems, for 1 ton of microalgal biomass there is a need of 200 tons of water. In contrast, for attached cultivation systems, for the same amount of biomass, 17 tons are required for water circulation which 4 tons of them are used for maintaining surface humidity in appropriate levels (Katarzyna et al., 2015). 2. While the high building cost of ordinary cultivation systems remains a serious problem for microalgal cultivation, this problem could be eliminated with biofilms. In the second case, the supporting surface of microalgal cells attachment could be cheap, easily available and reusable (Garbowski et al., 2017). 3. The biomass productivities of microalgal biofilms are much greater than those of suspended cultivation systems (Berner et al., 2015; Gao et al., 2015). Gross and Wen (2014) found that their microalgal biofilm system performance gave in average 302% biomass productivity as compared to the same performance in suspended cultivation system. 4. Microalgae attached to substrata seem to have high efficiency in wastewater treatment (Shen et al., 2018; Ma et al., 2018; Palma et al., 2017; Hodges et al., 2017). 5. In biofilms, microalgal cells are concentrated in a small footprint area (Choudhary et al., 2017). 6. While harvesting biomass is a nuisance for suspended cultures, this process is simplified with microalgal biofilms. In the second case, microalgal cells could be easily removed from the solid surfaces by scraping (Berner et al., 2015; Shen et al., 2014a; Zhang et al., 2018). In addition, the moisture content of biomass is very low that a further dewatering process is not necessary (Liu et al., 2013). The water content of Chlorella sp., harvested by scraping from an attached cultivation system was similar to the same species harvested by centrifugation from a suspended cultivation system under the same growth conditions (Johnson and Wen, 2010). After microalgal cells harvesting, the colonies which remain on the substrate will be the inoculums for the next growth cycle (Johnson and Wen, 2010). 7. Microalgae attached to the surfaces have better light availability as compared with those which are suspended in liquid medium (Katarzyna et al., 2015; Zhang et al., 2018). Lee et al. (2014) studied light penetration of attached and suspended culture systems. They concluded that the sunlight penetration on the upper layer of both culture systems was almost the same. However, the sunlight intensity

Culture system

Inoculum (g DW/L)

Culture medium

Petri–dish biofilm

1.8

BG11 medium

Supporting material

T (°C)

0.02

Glass fiber reinforced 26 plastic

Illumination CO2 (μmol·s−1·m−2) (% v/v) [light/dark circle (h)]

Duration (days)

Flow rate (L·min−1)

Rotation speed (rpm)

Species (Start exp)

150 [14/10]

14

–

–

Nannochloropsis N. oculata oculata

2

–

Glass

18–28

Wood biochar

20

18–28

–

Municipal WW

Algal biofilm membrane PBR (BMPBR)

40

Simulated secondary 0.28 effluent

Petri–dish biofilm system

20 ml Phosphate–enriched 0.01/Petri–dish Whatman GF/C filter 24 sub–culture tailing water

Flow–lane incubator

–

100 [16/8] blue – light 100 [16/8] green light 100 [16/8] yellow light 100 [16/8] red light 100 [16/8] purple light 100 [16/8] white light 100 [16/8] –

7

–

–

Shen et al., 2014a

13.5

Nylon mesh

–

Flexible fiber bundles 25–28

0.09

Polycarbonate slides

20

1.52 · 10

Mixed cellulose esters 26 membrane

4.76 · 10−2 5.80 · 10−2 20

–

–

18

–

–

4 8000 lx (maximum light intensity on the reactor wall) 134–203 [12/12] –

20

0.07

–

18

–

Random rotation

–

41

1.67

–

–

8

10−3

–

50 [14/10]

Hultberg et al., 2014

0.49 · 10−2

125 [16/8] −3

5.24 · 10−2

0.52 · 10−2

272–520 Natural – cond. (sunlight)

10 [16/8]

Chlorella vulgaris C. vulgaris, bacteria

0.81 · 10

85 [16/8]

Vertical algal biofilm (VAB)–enhanced raceway

3.38–3.67

−2

25 [16/8]

0.03 g wet Culture medium biomass·L−1 (2 g·L−1 urea) Culture medium (5 g·L−1 urea) – Synthetic WW with different nutrient load

Ref

5.32–15.76

Attached growth system

Membrane biofilm reactor

Biomass productivity (g·m−2·d−1)

1.77–3.87

1.8

Modified BG11 medium

Species (End exp)

Anabaena cylindrical Klebsormidium flaccidum Microalgae consortium (Chlorella sp., Pediastrum sp., Nitzschia sp., etc.) Chlorella vulgaris

A. cylindrical

110 mg DW

K. flaccidum

43.63 mg DW

–

9.10

Lee et al., 2014

C. vulgaris

0.05

Gao et al., 2015

Filamentous fungus, ≈0.64 Chlorella–like microalgae cells Bacteria, algae ≈0.11 cm−3·m−2·d−1 Algae ≈0.21 cm−3·m−2·d−1 Algae ≈0.06 cm−3·m−2·d−1 Algae, cyanobacteria ≈0.9 cm−3·m−2·d−1 Chlorella vulgaris C. vulgaris 9.27

Indigenous mixed microalgae Phototrophic biofilm material from WW treatment

Kholssi et al., 2018

Palma et al., 2017 Zippel and Neu, 2005

Rincon et al., 2017

12.64 0.16

Coral velvet

28

425 [14/10]

0.05

8

−3

(6.59–21.5)·10

–

Chlorella vulgaris, Oscillatoria

C. vulgaris, O. tenuis, S. obliquus, bacteria

10.54–14.68

Zhang et al., 2018

A. Mantzorou, F. Ververidis / Science of the Total Environment 651 (2019) 3187–3201

a. Permanently immersed biofilms Attached culture 0.3 Artificial seawater system (6–24 mM urea) Artificial seawater (6–24 mM nitrate) Artificial seawater (6–24 mM glycine) 4 Z8 medium Glass flask biofilm 10 cells ml−1 system

Cultivation area (m2)

3192

Table 3 Update of previous published table of experimental cultivation systems, based on microalgal biofilms. Species are referred to those inoculated and the beginning of the experiment (Start exp) and to those observed as predominant at the end of the experiment (End exp). Inoculum is expressed as dry weight of biomass per liter of culture medium (g DW/L). Illumination is expressed as μmol·s−1·m−2. Where it is possible, the hours of the light/dark cycle are also indicated. In some systems, where the biofilms are settled in a rotating area, the rotation speed is depicted in revolutions per minute (rpm). Otherwise it is not specified (–). Furthermore, (–) is used for data that are neither listed nor displayed in charts at source. Duration is referred to the time interval (days) between a growth (or regrowth) start up and a subsequent harvest. Biomass productivity is expressed as dry weight (g) of the biomass per unit of surface area, per day (g·m−2·d−1). Here, it is not specified if the mentioned biomass productivity refers to the initial growth or regrowth modes (subsequent growth cycles). Units are recorded where selected parameters were expressed differently. Some referred results have been calculated and extracted from graphs. Thus, these values are rough estimated (≈). avg. stands for average. Some parameters differ among the referred experimental setups and some others are not referred at all. That happens because of the different purpose of each experimental setup. Hence, we tried to refer the most appropriate combination. Cond. stands for conditions. WW: wastewater.

tenuis, Scenedesmus obliquus b. Biofilms alternated between gaseous and liquid phase Modified basal 0.02 Attached algal culture 106 cells·ml−1 medium system

0.1

Bench scale rotating – algal biofilm reactor (RABR)

8

19–23

756 [12/12]

–

60

0.01

2–5

WW effluent

0.19

290 [14/10]

–

26

–

4.8

WW effluent

2.72

Pilot Scale RABR

–

WW effluent

4.26

WW

Laboratory–scale rotating algal biofilm (RAB) system Laboratory–scale rotating algal biofilm (RAB) system

–

PVC

–

106 cells·ml−1

15

1.6

RABR–enhanced raceway pond

Attached algal culture system

–

Synthesized AMD

Cotton rope 14–24 Polypropylene Nylon Polyester Cotton (low thread) Cotton (high thread) Jute Acrylic Cotton cord 14–24

Rotating algae biofilm – reactor (RABR) Alga disk reactor 0.5

100 [24/0]

–

20

–

5.4

Cotton cord

9.6–19.2 208 avg. Natural – cond. (sunlight)

12

11.4

1.2

0.029

Braid cotton rope

20

230 [24/0]

–

84

–

–

M8a medium (30 mM urea)

0.06

Stainless steel mesh 37

5

7

0.54

9

Modified basal medium

0.02

Glass 26 fiber–reinforced plastic Plastic film Silicone film Maifan stone sheet Polyethylene foam Frosted glass Stainless steel sheet Polyurethane sheet Polycarbonate sheet Cotton duct 25 Cotton rag Cotton denim Cotton corduroy Cotton duct 25

399 [16/8] 399 [12/2] 12–635 [16/8] 399 [24/0] 399 [24/0] 100 [24/0]

10 –

15

–

8

4

Bold's basal medium

4.8 · 10−3

2

Bold's basal medium

45 · 10−3

Natural cond. (sunlight)

Scenedesmus dimorphus (UTEX 417) Chlorella protothecoides Chlorella vulgaris Scenedesmus obliqnus S. dimorphus (FACHB-496) Chlorococcum sp. Microbial consortium, dominated by Ulothrix sp. Mixed culture (microalgae, bacteria)

Mixed culture (microalgae, bacteria) Mixed culture (microalgae, bacteria) Polyculture microalgae Chlorella sorokiniana

Chlorococcum sp.

S. dimorphus (UTEX 417)

0.39

C. protothecoides

0.11

C. vulgaris

0.04

S. obliqnus

0.18

S. dimorphus (FACHB-496) Chlorococcum sp.

0.32

–

0.42

Orandi et al., 2012

–

≈2.12 0 0 ≈0.29 ≈1.42 ≈1.54

Christenson and Sims, 2012

0.53

≈1.62 ≈0.85 Diatoma sp., Pediastrum 20 sp., Chlorella sp. –

31

Filamentous cyanobacteria C. sorokiniana & bacteria

4.11

Chlorococcum sp.

110–120 [24/0]

–

15

–

30

Chlorella vulgaris

C. vulgaris

110–120 [24/0]

–

7

–

0.33 0.67 2 4

Chlorella vulgaris

C. vulgaris

Shen et al., 2014b

≈7.43 ≈6 ≈7.14 ≈11 ≈11 1.47

≈0.63 ≈1.1 ≈1.03 ≈0.95 ≈0.72 0.53 ≈0.6 ≈0.45 1.08 0.7 0.09 0.08 ≈2 ≈2.5 ≈2.5 ≈3.5

Christenson and Sims, 2012 Christenson and Sims, 2012 Hodges et al., 2017 Blanken et al., 2017a

Shen et al., 2014b

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Photo–rotating biological contactor (PRBC)

Stainless steel sheet 26

Gross et al., 2013

Gross et al., 2013

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(continued on next page)

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Table 3 (continued) Culture system

Inoculum (g DW/L)

Culture medium

Cultivation area (m2)

Supporting material

T (°C)

Illumination CO2 (μmol·s−1·m−2) (% v/v) [light/dark circle (h)]

Duration (days)

Flow rate (L·min−1)

Laboratory–scale rotating algal biofilm (RAB) system Pilot–scale RAB enhanced raceway pond system

2

Bold's basal medium

45 · 10−3

Cotton duct

25

110–120 [24/0]

7

–

6 4

–

Bold's basal medium

14

Cotton duct

January–February –

–

Rotating biological contactor (RBC)

–

M8a medium

M8a medium

0.36

0.36

Stainless steel woven rough mesh Stainless steel woven smooth mesh Sanded polycarbonate Stainless steel woven rough mesh

c. Permeated biofilm systems Multi–layers PBR 106 cells·ml−1

7

6

Single layer vertical plate attached PBR

–

38

422 [24/0]

7 15 mol·m−3

6

20

110–120 [24/0]

–

10

–

Modified basal medium

Glass fiber reinforced plastic Polyethylene foam Glass fiber reinforced plastic Polyethylene foam Cellulose acetate/ nitrate membrane

26

100 [12/12]

–

20

3.33

BG11 medium

10−3

11

Biofilm PBR

0.5 g·L−1 fresh inoculum

C. vulgaris

Chlorella sorokiniana

25

100 [24/0]

2

8

–

3 Chlorella 6 sorokiniana 11 20 3 11 20 3 11 20 15–60 Chlorella sp. tips·min−1

–

–

BG11 medium

Attached PBR with – multiple glass plates

Chlorella vulgaris

≈2.2 ≈3.5 ≈3 ≈3 ≈3.2 1.99

Gross et al., 2013

Gross et al., 2013

C. sorokiniana

20.7

Blanken et al., 2014

14.8

Polystyrene foam Cardboard Polyethylene fabric Loofah sponge

WW

C. vulgaris

Ref

18

Dairy manure WW 13.6 · 10−3

1.23

Chlorella vulgaris

Biomass productivity (g·m−2·d−1)

4.29

0.06 mol·m−3 0.5

Species (End exp)

May–June

0.7 mol·m−3

Attached culture system

Species (Start exp)

Artificial seawater enriched with full strength BG11 nutrients Artificial seawater enriched with f/2 nutrients BG11 medium

6 · 10−3

Cellulose acetate/nitrate membrane

30

BG11 medium

0.28

Concrete slab

25

12.8 [24/0] 14.2 [24/0] 18.7 [24/0] 29.3 [24/0] 56.2 [24/0] 134.5 [24/0] 55 [24/0]

C. sorokiniana

Chlorella sp.

Botryococcus braunii

B. braunii

B. braunii

B. braunii, other microorganisms

Scenedesmus S. obliquus obliquus Botryococcus B. braunii braunii Nanochloropsis Nanochloropsis OZ–1 OZ–1

18.7 19.5 20.1 18.5 ≈1.8 ≈2 ≈1.9 ≈1.8 ≈2 ≈1.9 2.57 1.47 0.58 1.28

3.19

2.71 10.46

≈3.5 ≈4 ≈4 ≈5.6 ≈7.8 ≈12 0.71

–

Scenedesmus obliquus

S. obliquus

–

35

0.15

–

Botryococcus braunii

B. braunii

Liu et al., 2013

≈5.25

≈5.25

–

Shen et al., 2015

≈6.8

C. fusiformis

9

Johnson and Wen, 2010

≈1.65 2.91

Cylindrotheca fusiformis 2

Blanken et al., 2014

Liu et al., 2013

Ozkan et al., 2012

A. Mantzorou, F. Ververidis / Science of the Total Environment 651 (2019) 3187–3201

Algadisk

–

0.03 1 2 3 22.4 avg. 261 avg. Natural – cond. (sunlight) 25.5 avg. 642 avg. Natural cond. (sunlight) 38 422 [24/0] 15 mol·m−3

Rotation speed (rpm)

PBR system semi–continuous flat plate parallel horizontal Phototrophic biofilm pilot–scale reactor

0.2–0.3

Vertical phototrophic biofilm reactor

Glass

–

Municipal WW effluents

8.08

Polyethylene–based Outdoor Natural cond. woven geotextile cond. (sunlight)

–

–

Synthetic WW

0.13

Polyethylene–based 21 woven geotextile

BG11 medium

0.09

Domestic WW

0.5

17 200 Flashing Cellulose light acetate/nitrate membrane Polycarbonate sheet 15–21.7 113–213 Natural cond. (sunlight)

Single layer vertical – plate attached PBR Attached PBR with multiple glass plates

Modified Chu 13 medium Chu 13 medium

0.08

Rotary biofilm reactor – (BIOALGA reactor) Capillary–effect–based 15 g·m−2 biofilm reactor (CBR)

Synthetic with 2% natural sewage BG11 medium

Biofilm reactor 9g cultivation chamber DW·m−2 Parallel plate reactor

Tubular periphyton bioreactor

–

–

Nitrate cellulose/cellulose acetate filter membrane

26

25

0.03/glass plate (not identifying the number of glass plates) 0.12 Polyurethane band

27–30

1.6

30

Lake water with 0.02 Woods Hole culture medium

Porous canvas

PVC

80 [16/8]

180 [24/0].

100 [24/0]

2

14

–

Scenedesmus S. obliquus, bacteria, obliquus other microbes Nitzschia palea N. palea, bacteria, other microbes Phototrophic Phormidium autumnale, biofilm Scenedesmus material from brasiliensis, Cymbella WW treatment minuta plant Microalgal – biofilm material from municipal WW treatment plant Scenedesmus S. dimorphus dimorphus

24

–

20 10.5 mg·L−1 (avg.) inorganic carbon

0.18

–

1

3

5.06 · 10−4

–

–

130

1.4

–

Scenedesmus obliquus

1

10

5.5 · 10−4

–

Botryococcus braunii

June–October

500 [24/0]

30

23.3 · 10−3

4.6–6 kW·m−2 – [11/13] 700 [12/12] 2.5 812 [12/12] 938 [12/12] 1134 [12/12] 938 [24/0] 938 [16/8] 938 [12/12] 938 [8/16] 700 [24/0] for 2 days & 1134 [8/16] for 6 days 700 [24/0] for 2 days & 1134 [8/16] for 2 days & 938 [16/8] for 4 days 2500 lx [14/10] –

Phormidium sp., Oscillatoria sp., diatoms, S. obliquus, bacteria B. braunii

0.01

2.1

Schnurr et al., 2013

2.8 2.7–4.5

Boelee et al., 2014a

2.7

Boelee et al., 2014b

9.13

Toninelli et al., 2016

1.92

Zamalloa et al., 2013

5.5–6.5

Cheng et al., 2013

49.1

20

–

2

6

–

–

Scenedesmus obliquus Desmodesmus sp.

8

–

–

Desmodesmus sp.

≈21.12 ≈21.85 ≈17.86 ≈23.24 ≈31.54 33.92 ≈16.83 ≈23.67 30.21

Travieso et al., 2002 Shen et al., 2018

27.95

4.5

0.02 · 10−3

–

Green microalgae, diatoms, Cyanobacteria bacteria, fungi, protozoa

Green microalgae, – diatoms, bacteria (such as Gammaproteobacteria, fungi, protozoa

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Unsterile secondary 0.14 WW effluent with synthetic medium

Ma et al., 2018

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on the bottom of suspended cultivation system was much lower than that of the biofilm systems. 8. As compared to suspended cultivation systems, biofilm systems can carry out during short hydraulic retention times (Boelee et al., 2014b). 9. The control of the cell growth area becomes easier when biofilm systems are set up in lagoons or in the ocean because of the high concentration of the attached cells on the substratum (Shen et al., 2018). Despite the beneficial aspects of microalgal biofilms, the ongoing research focuses on the potential drawbacks which they may have (Katarzyna et al., 2015). Those are referred below. 1. It is possible for the biomass to be detached from the solid surface. This may lead to the sedimentation and the loss of pollutants, which might had been bound to the biomass, toward the aquatic environment (Garbowski et al., 2017). 2. As mentioned above, microalgal biofilms through microbial proceeds, cause damage of metal surfaces such as erosion and decolorization (Katarzyna et al., 2015). Until now, a few microalgal biofilm systems have been developed (Table 3). Their design and geometric modulation have been chosen according to their purpose (Choudhary et al., 2017). However, these systems have been used mainly at a laboratory scale. Most of them concern the microalgal productivity as compared to the suspended cultivation systems. Some of them concern the production of microalgae with the aim of treating sewage from various pollutants (Katarzyna et al., 2015; Schnurr et al., 2013). A few of these efforts are referred below (see section 5). Nitrogen and phosphorus removal was greater with polyculture microalgal biofilm in a rotating algae biofilm reactor (RABR) system as compared to the open pond lagoon (Hodges et al., 2017). Also, BMPBR had better nitrogen removal capacity than suspended growth membrane photobioreactor (MPBR) did (Gao et al., 2015). Although, most

studies which are concerned microalgal biofilms are related to nitrogen and phosphorus removal, their application in wastewater treatment could be considered as a promising phycoremediation technology with the extra benefit of biomass manufacturing production for extra bioproducts use (Kesaano and Sims, 2014; Mantzorou et al., 2018). Ma et al. (2018) showed that the periphytic biofilm cultured in different Cu concentrations (2 and 10 mg·L−1), had high removal efficiency in Cu (99%). This is due to the EPS production which offers a great number of functional groups for the metal complexation (Palma et al., 2017). Christenson and Sims (2012) showed the significant potential of a laboratory–scale photorotating biological contactor (PRBC), established with a microbial consortium dominated by Ulothrix sp., to remove 20–50% of toxic metals (such as Cu, Ni, Mn, Zn etc.) from synthesized acid mine drainage (AMD). Nevertheless, such systems in wastewater treatment still remain manly in experimental stage (Kesaano and Sims, 2014). The design and the large-scale operation of such systems become difficult because they are not fully investigated in the so far literature (Kesaano and Sims, 2014). Other systems concern the ability of microalgae to produce biofuels (Ozkan et al., 2012). Some of these constructions are described below. 5. Main microalgal based biofilm cultivation systems Based on the liquid medium supply and the biofilm arrangement, microalgal biofilm designs were first divided by Berner et al. (2015) to: i) constantly submerged systems, ii) intermittently submerged systems and iii) perfused systems. According to his classification, these systems could be categorized into: i) permanently immersed biofilm i.e. those which are permanently immersed in the liquid culture medium, ii) biofilms between two phases i.e. those which are alternated between gaseous and liquid phases and iii) permeated biofilm systems i.e. those in which culture medium is provided directly to the substratum (Fig. 2). The last categorization could be considered as more descriptive

Fig. 2. Simplified designs of some microalgal biofilm systems: [a] System of permanently immersed biofilms; [b] and [c] biofilms between two phases; [d] permeated biofilm system.

A. Mantzorou, F. Ververidis / Science of the Total Environment 651 (2019) 3187–3201

as far as microalgae and medium arrangement are concerned. In Table 3, some experimental cultivation systems based on microalgal biofilms are described with respect to their selected parameters. 5.1. Permanently immersed biofilms A simple construction was designed by Shen et al. (2014a) to improve the lipid yield of Nannochloropsis oculata. In 500 ml beakers, 4 layers of glass fiber–reinforced plastic were placed as a supporting material. The beakers were filled with a suitable liquid culture medium so as to cover the upper supporting material as well. To examine oil production, three parameters were tested: nitrogen deficiency, high illumination and the combination of the two previous parameters (Table 3a). In the study of Lee et al. (2014) mesh–type materials were placed in open pond in order to evaluate the biofilm formation of a microalgae consortium (such as Chlorella, Nitzschia, Scenedesmus etc.). Their performance was compared to another one, where microalgal species were suspended in the same open pond (Table 3a). An enclosed tubular biofilm PBR was presented by de Godos et al. (2009). This system was inoculated with the microalga Chlorella sorokiniana and a mixed bacterial culture. The system consisted of a 14 m transparent PVC tube, arranged horizontally in a coil manner. Pretreated (centrifuged) swine slurry loading was provided to the system. The system also included a closed storage reservoir for effluent removal and culture recirculation. The enclosed tubular biofilm PBR was 3− evaluated based on the C, NH+ 4 , and PO4 removal from pretreated piggery wastewater. 5.2. Biofilms between two phases A rotating algal biofilm (RAB) was designed by Gross et al. (2013). In a plexiglass chamber, placed on a rocker shaker, various materials were tested for their ability as adhesion materials of Chlorella vulgaris cells (Table 3b). This system was rotating so that when one corner of the triangle was submerged in the liquid medium, the other two corners were exposed to atmospheric conditions. In this way, microalgal biofilm was exposed both to atmospheric CO2 and nutrients as well. This microalga was tested for its biomass yield, biomass productivity and its chemical composition. The RAB system performance was compared with the same performance at a flat panel PBR. Another concept was introduced by Johnson and Wen (2010). In their effort, Chlorella sp. was left to be attached in different supporting materials (Table 3b). Their system was composed of a growth chamber in which the tested materials were placed in. The chamber was gently moved with the help of a rocking shaker. As the shaker was moving the chamber, the latter was submerged by half so that when its half was sinked on the liquid medium, the other half was exposed to illumination. Their system was compared with suspended culture which took place on the same chamber without any supporting material. A PRBC was designed by Orandi et al. (2012). For the biofilm establishment, a microbial consortium, dominated by green microalgae (Ulothrix sp.) was used (Table 3b). For the construction of the PRBC, 16 polyvinyl chloride disks with roughened surfaces were used for the biofilm development. The disks placed in a shaft, were submerged by 40% in a plexiglass tank. The shaft was connecting with a motor for controlling the rotational speed of the disks. The plexiglass tank was coupled to a feed container for providing the liquid media (multi–ion synthesized simulated AMD) to the disks. Blanken et al. (2014) described a PBR devise based on a rotating biological contactor (RBC). They used Chlorella sorokiniana for evaluating the potential of such design in terms of photosynthetic efficiency, productivity and cultivation stability. In detail, 4 discs were submerged (by 42%) inside a watertight container. The tested disc materials were two stainless steel woven meshes (of different tread thickness and particle pass size) and one sanded polycarbonate disk (Table 3b).

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5.3. Permeated biofilm systems A different biofilm design was described by Ozkan et al. (2012). Botryococcus braunii was cultivated axenically in a biofilm PBR which was composed of a growth surface (thick concrete layer) over a wood support plate (Table 3c). The nutrient flow (which was provided from dripping nozzles above the growth surface) was achieved by growth surface tilt. Liu et al. (2013) introduced 2 types of attached cultivation bioreactors (Table 3c). The first one which they called it ‘single layer vertical plate’ was composed of a glass plate, embedded in a glass chamber. While the one plate surface was illuminated, the other surface was covered with a filter paper, in which the microalgal cells were attached. The second type of attached cultivation bioreactor was similar to the first. The only difference was that in the second bioreactor, the glass chamber was composed of more than one, glass plates. The glass plates were placed in a way that each of them received light of different intensity. In this research Scenedesmus obliquus, Botryococcus braunii, Nanochloropsis OZ-1 and Cylindrotheca fusiformis were involved. The culture medium was provided by dripping down to the interface of the glass plate and the filter paper. Schnurr et al. (2013) described a semi–continuous system consisting of 12 rectangular flat plates placed in parallel, where the adhesion of Scenedesmus obliquus and Nitzschia palea cells was performed on a glass substrate (Table 3c). This system was designed for studying neutral lipid productivities under nitrogen and silicon starvation and growth kinetics. In a similar manner, but using filter paper instead of glass, as adhesion material, Cheng et al. (2013) examined the content of Botryococcus braunii in lipid and hydrocarbons when the microalga was subjected to nitrogen sufficiency and deficiency. In both cases, culture medium was provided by flowing through the substrate (Table 3c). 6. Factors affecting microalgal biofilms The formation and structure of biofilms is contingent on various parameters and interactions among living organisms that conduce to settlement and growth (Bott, 2011). In opposition to bacterial biofilms, there is no much information about the factors affecting microalgal biofilms (Irving and Allen, 2011). The growth optimization under various environmental parameters still remains a big challenge. This is a bottleneck in designing microalgal biofilms for full–scale operation especially in wastewater treatment (Kesaano and Sims, 2014). Nevertheless, a thorough understanding of the most influential parameters is crucial for the technical development and the design of microalgal biofilms. These information will provide the tools for the control of such systems and the survival of microorganisms which participate in (Choudhary et al., 2017). 6.1. Selection of proper microalgal strain The choice of the proper strain is probably the most crucial factor in biofilm formation. Microalgal species have different properties and behaviors. Thus, some strains grow better in solid surfaces while others are favored suspended in liquid media (Katarzyna et al., 2015). As compared with Scenedesmus obliquus, Nitzschia palea has been proved to be a very adherent microalgal species with high biomass productivities and robust biofilm forms (Schnurr et al., 2013) (Table 3c). Furthermore, some strains cannot form single species biofilm, but they should be assisted by other adherent microorganisms (Li et al., 2007). Irving and Allen (2011) showed that in axenic conditions, between Chlorella vulgaris and Scenedesmus obliquus, the former has lower initial attachment (expressed as cells·cm−2) than the latter did, when microalgae left to be attached in various materials coupons. It is also a fact that the attachment mechanism for each organism group differ from one another. The attachment of most diatoms proceeds with the EPS production in the form of apical pads, stalks, cell coatings and mucilage pads

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while the attachment of filamentous green algae proceed mainly by the form of holdfast. Most cyanobacteria attach to the substrata via the EPS production in a similar form to diatoms and bacteria (Sekar et al., 2004). 6.2. Nutrients Nutrient availability is crucial for the microalgal biofilm formation (Katarzyna et al., 2015). The biofilm growth pattern is characterized from nutrient removal trends. At the beginning of the growth phase, the nutrient uptake efficiency is relatively low due to inefficient settlement and adjustment of the biofilm cells. As growth soars, nutrient uptake increases. Finally, at the death phase, this efficiency is reduced as microalgal biofilm loses its integrity because of sloughing (Boelee et al., 2011; Kesaano and Sims, 2014). As microalgal biofilms could be considered as different life forms from suspended microalgal cultures, there seems to be differences in responding to nutrient uptakes and deficiencies as compared with suspended cultures (Fields et al., 2014; Shen et al., 2014a). This was proved by Schnurr et al. (2013) who studied the potential of Scenedesmus obliquus and Nitzschia palea, grown in a biofilm system, for lipid stimulation under nutrient starvation (Table 3c). They concluded that in contrast to suspended growth mode, nutrient depletion did not favor lipid accumulation to microalgae. Also, it has been shown that increasing nitrogen concentration results in elevated EPS production from green algae species and diatoms (Schnurr and Allen, 2015; Shen et al., 2015). Furthermore, elevated concentrations of nitrogen and phosphorus in the surround medium lead to notably higher photosynthetic biomass accumulations in a biofilm structure as compared to bacteria (Kebede-westhead et al., 2003; Villanueva et al., 2011). Under nutrient starvation, EPSs could be degradated in order for the nutrients enclosed to the matrix, to be used for microalgae feeding or for the microalgae detaching and repopulation in better nutrient existing conditions (Schnurr et al., 2013). 6.3. Light availability In opposition to bacteria biofilms, microalgal biofilms are strong depended by light which is the main energy source for their development (Zippel and Neu, 2005). By a light saturation point (200–400 μmol·m−2·s−1), microalgal growth increases in a linear way with increasing light (Boelee et al., 2014a) (Table 3c). In spite of bioreactor configuration or cultivation design, some authors consider light to be the key factor for microalgal growth optimization (Toninelli et al., 2016). Not only is the adhesion of microalgae influenced by light availability but the synthesis and the accumulation of EPSs as well (Katarzyna et al., 2015; Zippel et al., 2007). Nevertheless, too much light in the upper biofilm layers (photoinhibition) or little light in shaded layers (photolimitation) can inhibit microalgal biofilm growth (Kesaano and Sims, 2014). Concerning lower biofilm layers, in light limited conditions (because of too thick biofilm or insufficient light intensities), bacteria, EPSs and other non–photoautotrophic materials would be dominated (Schnurr and Allen, 2015). It is well understood that the microalgal growth is increased as the cells are closed to the light source. This was proved by Liu et al. (2013) who examined the growth of Scenedesmus obliquus in an attached PBR system (Table 3c). They evaluated the biomass concentrations at various distances of the cultivation surface from the light source. They concluded that the light dimming caused a decreased growth rate of the microalga. Furthermore, Hill and Larsen (2005) showed a taxinomic difference among shaded, unshaded and UV biofilms. The microalgal composition of shaded biofilm was different from those of unshaded and UV biofilms. Another parameter was tested by Hultberg et al. (2014) who studied the effects of different light colors on biofilm formation by Chlorella vulgaris. They found that white, blue and purple light resulted in sooner and higher biofilm formation as compared to the treatments with red, yellow and green light.

Changes in light regime could influence the nutrient uptake by microorganisms. This was confirmed by Sukačová et al. (2015) who examined the phosphorus uptake from a biofilm composed of a microalgae consortium. During a continuous light regime, the efficiency of phosphorus removal reached 97% while the phosphorus uptake ranged between 36 and 41% during day–night light regime and using solar radiation. (Hultberg et al., 2014). 6.4. Temperature Temperature is another factor which could affect species composition, microalgal growth rates and grazing activity (Kesaano and Sims, 2014). Increasing temperature to a point results to a high metabolic activity and therefore to elevated biomass productivity (Gross and Wen, 2014). Also, the enzymatic activities by which organic matter is degraded by bacteria, are enhanced by increasing temperature (Choudhary et al., 2017). Nevertheless, each microalgal species has different temperature requirements. Generally, concerning mesophilic microalgae, 20–25 °C is the optimum temperature for maximum growth rates (Boelee et al., 2014a). Microalgal biofilm systems are more susceptible to temperature fluctuations than suspended culture systems because of their considerable low water quantities as compared to the latter. In the last case, water have a buffer role on temperature (Ozkan et al., 2012). Furthermore, the water conservation is crucial due to potential evaporation losses by high temperatures. It has been calculated that evaporation losses of a biofilm photobioreactor (BPBR) (with characteristic length 10 m, water layer thickness 0.05, biofilm layer thickness 0.005 and water velocity 0.05 m·s−1) in autumn, winter, spring and summer could reach 3.4, 1.0, 6.0 and 7.3 L·m−2·d−1 respectively (Murphy and Berberoğlu, 2011; Kesaano and Sims, 2014). 6.5. pH Microalgal biofilms depend on pH. Biofilm, as a new micro– environment, has different pH from the surrounding medium. pH changes are observed among various biofilm layers (Katarzyna et al., 2015). Also, pH is crucial for the cell establishment (Sekar et al., 2004). EPSs and microalgal membrane are composed mostly of proteins, polysaccharides and lipids. Different pH could influence the components of extracellular metabolites. At low pH values, the dissociation of amine and carboxyl groups is enhanced and inhibited respectively. This leads to the weakening of negative surface charge of microalgae and subsequently to increase of microalgal adhesion. Generally, microalgal growth is favored from pH between 6 and 9 (Shen et al., 2014b). Sekar et al. (2004) who examined the effect of pH on microalgae adhesion showed that Nitzschia amphibia had significantly better attachment at pH ≥ 7. 6.6. CO2 Carbon is essential to photosynthetic microalgae for various metabolisms (Gross et al., 2013). Carbon is afforded to them from CO2(aq) and bicarbonate (HCO− 3 ) in wastewater. Atmospheric CO2 and this resulting from organic carbon degradation by bacteria, are also available to the microalgae (Kesaano and Sims, 2014). Nevertheless, concerning phototrophic growth, CO2 is considered to be the primary carbon source. If the CO2 consumption is more than CO2 supply, the microalgal productivity will be diminished because of carbon limitation (Blanken et al., 2014). Concerning suspended culture systems, models have been constructed with which CO2 loss could be minimal through the mass transfer from the gas to the liquid phase. On the other hand, while high CO2 levels lead to high productivities, systems which could confine CO2 losses have not been thoroughly developed for microalgal biofilms (Blanken et al., 2017b). However, some systems design (such as RAB

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system) are exposed to the gas phase, and permit to microalgal biofilm to be exposed directly to the gaseous CO2 instead of dissolving CO2 in the medium. The increase of CO2 concentration in the inlet air of RAB, beyond the atmospheric levels, did not increase more the biomass productivity of Chlorella vulgaris, maybe because of the efficient transfer of gaseous CO2 to the microalgal cells (Gross et al., 2013). 6.7. Attaching material Various materials for the microalgal attachment have been tested and compared. Among the materials used for this purpose are glass, polystyrene foam, muslin cheesecloth, polyurethane foam, vermiculite, jute, polyester, cardboard, poly–lactic acid, fiberglass, cotton duct, cotton rope etc. (Gross et al., 2013; Johnson and Wen, 2010; Shen et al., 2014a). The evaluation of these materials is usually based on their durability and reusability as well as their cost and the degree of cell attachment (Gross et al., 2013). However, to date, there are no standard materials proposed as biofilm development substrata (Kesaano and Sims, 2014). Biofilm formation and growth is affected differently by miscellaneous materials. The properties of them are very important for understanding and predict the biofilm development (Schnurr and Allen, 2015). It has been proven that the texture and roughness of the supporting materials play a key role in cell adhesion (Katarzyna et al., 2015). In general terms and with only a few exception results, the cell adhesion is favored of hydrophobic materials (Zeriouh et al., 2017). However, there are some who argue that hydrophobicity does not play any role in microalgae adhesion. Irving and Allen (2011) showed that the substrate hydrophobicity (by means of contact angle) has not been correlated with microalgae colonization. Gross et al. (2013) tested a total of 16 materials for attaching Chlorella vulgaris in a RAB system. Some of them are referred in Table 3b. Cotton duct was proved to be the most suitable material because of its durability, low cost and good cell attachment. They attribute the suitability of this material to its crevices, which protect the cells from the sheer force, when they are moving through the liquid. In addition, among loofah sponge, cardboard, polyethylene landscape fabric, polystyrene foam, nylon sponge and polyurethane foam, polystyrene foam was proved to be the most suitable material for the biofilm formation by Chlorella sp., in terms of physical robustness, biomass harvest, microalgal growth and substratum reusability (Johnson and Wen, 2010) (Table 3b). 6.8. Flow velocity The biofilm formation is strongly depended on the flow of liquid phase, in which the biofilm is immersed and supply the microorganisms with nutrients (Choudhary et al., 2017). The liquid must to circulate with an adequate velocity in order to supply microorganisms with nutrients and to remove waste products. However, high velocity could cause shear stress on biofilm (Berner et al., 2015; Choudhary et al., 2017). Furthermore, turbulent flows of liquid medium can cause cell detachment and consequently decrease of biofilm thickness (Berner et al., 2015; Katarzyna et al., 2015). Johnson and Wen (2010) noticed that when their attached cultivation system kept stationary, the microalgal cells settled on the supporting material while they could not be firmly attached. Nevertheless, at the beginning of cultivation, cell attachment is favored by a low flow velocity while a higher flow is adequate thereafter in order for the microalgae to be provided with nutrients for their further growth (Berner et al., 2015). In some configurations, where the microalgal biofilms are settled in a rotating area alternating them between gaseous and liquid phase, the rotational speed is crucial for the cell attachment. A proper speed protects microalgal biofilm from shear stress which influences the cell attachment and biofilm thickness (Gross et al., 2013). Johnson and Wen (2010) reported that when their attached culture system was kept stationary, the cells of Chlorella sp. settled on the substratum without

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creating a firm biofilm structure. They concluded that the settlement of some algae which are naturally found in aquatic environments with high current velocities, are favored of rocking which mimics surge or wave effect. The best rotational speed for biofilm establishment in a PRBCs design has proven to be between 1 and 10 rpm (Orandi et al., 2012). Nevertheless, Blanken et al. (2014) who examined the influence of disk rotation speeds (3, 6, 11 and 20 rpm) of a RBC based PBR design, on Chlorella sorokiniana growth found that the effect on biofilm productivities were minimal (Table 3b). Gross et al. (2013) found that in a low rotation speed (b4 rpm) the microalgal biofilm was exposed to the gaseous phase too long and tended to lose its wetness. On the other hand, at high rotation speed (6 rpm) the biofilm sheared off because of high shear stress (Table 3b). 6.9. Presence of other microorganisms Until recently, bacteria were being considered as contamination factor for microalgal cultures. The last few years this fact has changed. Recent studies have proved microalgae–bacteria relationship to be essential in algal biotechnology since it favors microalgal growth and flocculation procedure (Fuentes et al., 2016). Bacteria which are present in wastewater have been proven to create a favorable environment on which the microalgal biofilm is developed (Schnurr et al., 2013). It is well known that microalgae provide O2 which is used by bacteria for the oxidation of NH4 and organic matter. On the other hand, CO2 is provided by bacterial respiration for the microalgal photosynthesis (de Godos et al., 2009). It has been confirmed that in the initial colonization process, bacteria strongly interact with microalgae in the biofilm structure. It has been shown that the biofilm formation in bacteria is associated with Toxin–Antitoxin Systems (Karimi et al., 2015). These systems contribute to the processes associated with controlling bacterial survival (Wen et al., 2014). As bacterial abundance and diversity is higher, more carbon sources are available to microalgae and many more microalgal cells are attached to the solid surfaces (Choudhary et al., 2017). The thickness of Chlorella vulgaris biofilm under non–sterile conditions was much greater than those under sterile conditions. On the other hand, Scenedesmus obliquus formed biofilms with comparable thickness in both sterile and non–sterile conditions. Nevertheless, in both cases of microalgae, nor S. obliquus or C. vulgaris managed to dominate in non–sterile conditions (Irving and Allen, 2011). Furthermore, Rivas et al. (2010) showed the enhanced and reduced growth of Botryococcus braunii biofilms under the presence of Rhizobium sp. and Acinetobacter sp. respectively. Also, it has been observed that epiphytic bacteria, isolated from Ulva lactuca and Utricularia reticulata inhibited the growth of Nitzschia paleacea and Cylindrotheca fusiformis. Furthermore, the cells of Chattonella antiqua and Eucampia zodiacus were lysed by Alteromonas strains (Mieszkin et al., 2013). Nonetheless, Kawamura et al. (1988) demonstrated that bacteria have no necessarily effect on some microalgae attachment. This was proved by Alcaligenes sp., which has no influence on the attachment of Synedra sp. to substrata. 7. Conclusions Microalgal biofilms is an active research field in early stage. The experimental results have been characterized quite promising. These systems, applied mainly in laboratory scale, offer valuable information but some points between the factors and the way that influence the microalgal biofilm development remain ambiguous. Although, some efforts have been done, it is necessary such systems to operate in larger scale and real conditions for their actual assessment. Different chosen variables hamper the assessing of such system performance. For evaluating of their potential, it is necessary to be assessed using the same baselines of construction and operation.

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