Apparatus for microalgal or cyanobacteria growth, methods and uses thereof

Abstract

The present disclosure relates to an apparatus for microalgae and / or cyanobacteria cell culture comprising a unitary body formed from a hydrophobic material; and two or more microcapillary channels extending along the unitary body; wherein the inner surface of the microcapillary channels is coated with nutrients suitable for microalgae and / or cyanobacteria cell culture; and wherein the hydrophobic material is transparent to visible light, and to a kit and sensor thereof. A method for culturing microalgae and / or cyanobacteria in an apparatus comprising a unitary body formed from a hydrophobic material and two or more microcapillary channels extending along the unitary body, wherein the hydrophobic material is transparent to visible light is also disclosed.

Description

D E S C R I P T I O NAPPARATUS FOR MICROALGAL OR CYANOBACTERIA GROWTH, METHODS AND USES THEREOFTECHNICAL FIELD

[0001] The present disclosure relates to an apparatus for microalgal and / or cyanobacteria cell sensing and culture, and to a methodology of micro-cultivation of microalgae based on the disclosed apparatus. BACKGROUND

[0002] Microalgal cultivation is increasingly important for industrial, agricultural, and environmental monitoring. Current methodologies for monitoring microalgal growth require periodical sampling, bulky laboratory equipment, and specialized personnel.

[0003] Microalgae consists of photosynthetic microorganisms mostly found in aquatic habitats including lakes, ponds, rivers and oceans. These microorganisms are currently viewed as one of the most interesting sources of biofuels, bioactive compounds for medicinal purposes and nutrition applications with a well-developed worldwide industry already dedicated to biomass accumulation, harvest and downstream refinery of the active compounds [1], [2]

[0004] Equally, microalgae can cause severe environmental problems, for instance eutrophication by depleting oxygen levels in surface water systems or contaminating drinking water reservoir with toxins. Indeed, some species bloom in still reservoirs, especially when enough light, temperature and nutrients are available. Harmful Algal Blooms (HABs) occur when the blooming species produce toxins such as geosmin and microcystin, which cause human and animal hazards. The most impacted industries include drinking water, wastewater, recreational lakes, agriculture, aquaculture and the power generation units. Thus, regular monitoring of microalgal growth is therefore essential for a range of biotechnological applications and environmental water surveillance [3].

[0005] However, this currently relies on conventional microbiological techniques, which are time consuming, require regular field sampling and access to microbiological labs with sterile conditions and skilled personnel [4]. Previous attempts to miniaturise and automate microalgal cultivation yielded complex bioreactor designs and limited growth rates [5].

[0006] In laboratory settings, microalgal growth curves are generally determined based on counting and usually start with an initial lag phase, followed by exponential, linear and a declining stage [6]. This is carried out in cultures grown in lab-scale tubes and Erlenmeyer flasks. In species with high ecologicaland biotechnological importance such as Chlorella and related species [7], growth is determined in more sophisticated bioreactors.

[0007] Microfluidic devices offer the opportunity to automate and make it portable the cultivation of microalgae. Previously, several complex devices have been proposed and are reviewed in detail in [5], some of those relying on the integration of microfluidics with bioanalytical features such as determination of lipid content [8]. All those concepts made use of optical properties and advanced features in microfluidic devices and operated as continuous-flow microscale photobioreactors [9]. The generic components of microfluidic systems applied to cells includes microchannels, valves, pumps, growth chambers and the perfusion of nutrients or molecules in study [8].

[0008] Kim et al. [9] describes the application of microfluidic lab-on-a-chip systems for microalgal biotechnology, especially microalgae-based biofuels, including microsystems for single-cell resolution high-throughput cell identification and separation, highly efficient cell transformation, high-throughput parallel cell cultivation, cell harvesting, and cell analysis applications. The disclosed systems require continuous growth under continuous flow conditions or in encapsulated droplet – which also involves continuous flow. Thus, this document is silent on sealed lab-on-a-chip systems, where the growth of microalgae occurs in a static self-enclosed unitary body.

[0009] Bodenes et al. [5] teaches that cultivation of microalgae in microfluidic devices requires a design for trapping the cells and study microalgae at cell scale, or the same population undergoing a continuous media. Accordingly, the same study identified three categories of microreactors for cultivation of microalgae, being (1) mechanical traps; (2) droplets; and (3) microchambers, in all cases they involve trapping of cells and continuous supply of media.

[0010] Reis et al.

[0010] describes a mass-manufacturable dipstick, based on melt-extruded Teflon® fluorinated ethylene propylene (Teflon FEP) microfluidic film coated with poly(vinyl alcohol) (PVOH), operating based on capillary forces. This dipstick has applications in the detection of bacteria such as E. coli and antibiotic susceptibility testing

[0011] and blood testing. However, the application of said dipsticks to microalgae culture is challenging compared to bacteria due to microalgal need of suitable light intensity. Also, it is known from the state of the art that the cultivation of microalgae in microcapillaries requires supply of fresh media, mixing, or aeration[5], [8] .

[0011] Document EP3172565B1 relates to an assay device having a unitary body with an exterior surface, the unitary body being substantially transparent to visible light and formed from a material having a refractive index in the range 1.26 to 1.40, the refractive index being measured at 20 °C with light of wavelength 589 nm. The unitary body is formed from a hydrophobic material, and at least two capillary bores extending internally along the unitary body, wherein at least a portion of the surface of each capillary bore includes a hydrophilic layer for retaining an assay reagent, and wherein thehydrophilic layer is also substantially transparent to visible light to allow optical interrogation of the capillary bores through the capillary wall. However, use of said assay device on direct non-invasive optical monitoring of microalgae populations is not disclosed.

[0012] Document EP2550147B1 describes a device for carrying out an immunoassay, the device having: a unitary body with an exterior surface, and at least two capillary bores extending internally along the unitary body. The unitary body is substantially transparent to visible light to allow optical interrogation of the capillary bores, and the device is formed from a material having a refractive index which is in the range 1.26 to 1.40 and is within plus or minus 0.07 of the refractive index of the sample fluid, the refractive index being measured at 20 °C with a light of wavelength 589 nm. Regardless, this document is silent on the use of said device on the microcultivation of microalgae or cyanobacteria.

[0013] Currently, the state of the art involving HAB-related problems focus on methods for eliminating the algal bloom rather than avoiding it. The works that aim detection of microalgae prior to proliferative events, are based on the expensive, time-consuming sequencing methods for molecular identification of the blooming species.

[0014] These facts are disclosed in order to illustrate the technical problem addressed by the present disclosure.GENERAL DESCRIPTION

[0015] The present disclosure relates to an apparatus for microalgae and / or cyanobacteria cell culture, and to a methodology of short and long-term micro-cultivation and preservation of microalgal based on microcapillary strips. In particular, it describes a fluoropolymer (preferably fluorinated ethylene propylene) microcapillary strip for passive sampling, via capillary forces, and further micro-cultivation of microalgal cells.

[0016] In an embodiment, the microalgal cells are from Parachlorella kessleri (previously known as Chlorella kessleri). These cells provide a good optical contrast due to their bright green colour and spheric morphology without appendages such as flagella, spines and ornaments elements. In another embodiment, microalgal cells are from Lobochlamys segnis.

[0017] The disclosed microcapillary strips are an alternative to existing and conventional long term preservation methods such as cryobanks with continuous supply of liquid nitrogen, and long term and expensive cell recovery procedures, and handling by specialized personnel. The time a culture extracted from a cryobank to recover is around 1 month. When cultures are recovered, it is common to observe cell death due to cryo-preservation damage. Surprisingly, in the disclosed microcapillary strips, Parachlorella kessleri cells with no nutrient renewal and inside the disclosed microcapillaries can last atleast for 2 years, under low light conditions (15 µmol / m2 / s) and do not suffer from large recovery times, e.g. a culture is fully developed in around one week with no sub-selection / physical damage to cells.

[0018] In an embodiment, microcapillary strips can be prepared from a fluoropolymer selected from a list comprising fluorinated ethylene propylene (FEP), tetrafluoroethylene hexafluoropropylene vinylidene fluoride (THV), perfluoroalkoxy (PFA), polytetrafluoroethylene (PTFE), ethylene tetrafluoroethylene (ETFE), poly(chlorotrifluoroethylene) (PCTFE), polydimethylsiloxane (PDMS), and mixtures thereof.

[0019] In an embodiment, the excellent light properties and O2 / CO2permeation of Teflon FEP material, allowed to image and sustained growth of the microalgal cells over a period of 22 days. Surprisingly, the cultivation of microalgae in a static self-enclosed unitary body - the microcapillaries (microcapillary channels) - was feasible without the need of supply of fresh media, mixing, or aeration during the 22 days of growth, and with high growth rates. In a further embodiment, the strip may range in length from a few centimetres to several meters and can be configured to roll and stand laterally. This design allows the system to occupy only a few centimetres of bench space, even when the strip is several meters long, thereby enabling storage of a larger inoculum of the desired species.

[0020] In an embodiment, the use of the disclosed apparatus as micro-cultivation platform provides a portable, simple, cost-effective, and reliable system which allows growth of microalgae at the most favourable conditions.

[0021] In an embodiment, the disclosed apparatus for microalgae and / or cyanobacteria cell culture can be used in the early detection of harmful algal blooms, prediction of harmful algal blooms, or assessment of water quality, using static culture conditions. A capillary sensor comprising the disclosed apparatus is positioned at the surface of the water tank to be monitored, where phytoplankton (aquatic microalgae and cyanobacteria) lives. A water sample is collected to the apparatus and retained inside the microcapillary channels of the sensor. The captured cells remain alive and growing inside the sealed microcapillary channels. The cells proliferate inside the microcapillary channels up to one month and can be observed under a microscope or a smartphone.

[0022] In an embodiment, the microorganisms are then analysed with microscopy for timely detection of HAB forming species. This method to monitor and model algae growth is highly attractive to both industry and academia because it is low-cost and provides accurate and unprecedented readouts of cell growth dynamics and single cell morphology over time.

[0023] The disclosed capillary sensor has a low-cost installation and utilization, and also allows an optimal growth of photosynthetic microalgae and cyanobacteria species due to its transparency and circular shape. Even in low numbers, the cells are able to grow inside the microcapillary channels, which enables the early sensing of the toxic species presence and development / growth stage.

[0024] In an embodiment, the sample may be monitored with a microscope, or ultimately with a smartphone, inside the microcapillary channel. In a further embodiment, microscopic analysis can be performed directly on the microcapillary strip, by visual inspection of the cells and by performing taxonomic identification.

[0025] In an embodiment, the disclosed system permits monitoring and preventing HABs in both water reservoirs and in the environment, while allowing the investigation of growth and morphological changes of industry-relevant species, such as cyanobacteria.

[0026] The advantages of the mentioned technology include the ability to visually inspect and monitor microalgae and cyanobacteria growth in marine environments on-site or in-situ, either through a microscope or a smartphone, facilitating timely detection of growth stages to prevent harmful exponential algae growth and subsequent release of toxins into the water. Additionally, it allows for real-time studies of cyanobacteria growth and morphological changes, marking a significant advancement in research capabilities.

[0027] In an embodiment, the disclosed apparatus can be functionalized by covering its inner surface with specific growth nutrients, which favor the growth of the target species to be detected and studied. The microcapillary channels can be coated with a selection of nutrients specifically selected for the growth of the target species. This enables a more predictive approach for HAB. In a further embodiment, the nutrients are adherent to the inner surface of the microcapillary channels and serve as growth promotors of captured cells even in low numbers, at the start of proliferation.

[0028] In another embodiment, computed implemented methods can be used connected with the disclosed stick, for HAB predicting and cell determination.

[0029] The apparatus disclosed in the present disclosure allows the early detection of HABs, while providing improved growth of microalgae and cyanobacteria, as the same growth is achieved using batch conditions as compared to conditions. Also, the transparent microcapillary channels allow a real time observation of microalgae and cyanobacteria with a microscope, or ultimately with a smartphone camera.

[0030] The disclosed method is a fast, low-cost, discardable method for analysing water quality. It originates a timely information on the onset of a blooming event, therefore preventing all the harmful diseases caused by toxic water consumption by direct drinking or by eating contaminated vegetables or animals from aquaculture farms, as well as contaminated recreational lakes and water-related companies which struggle with the problems caused by blooming algae.

[0031] The apparatus may be functionalized by covering the inner surface with specific growth nutrients, which favor the growth of the target species to be detected and studied. The microcapillary channels can be internally coated with a selection of nutrients specifically selected for growth of targetHAB forming species. The nutrients are adherent to the inner surface and serve as growth promotors of captured cells even in low numbers, at the start of proliferation.

[0032] The disclosed micro-cultivation methodology based on microcapillary strips or ‘dip sticks’, which can be optionally functionalised, allows sampling and optical monitoring of microalgal growth without the need of complex, microfluidic design. Surprisingly, cultivation experiments carried out in static microcapillary strips, with no replacement of growing media nor physical aeration, showed a maximum growth rate of 0.37 day-1for P. kessleri, matching the value observed for a sparged Erlenmeyer, and representing a >3-fold improvement compared to batch unsparged Erlenmeyer (growth rates of 0.12 day-1). The good optical properties of fluoropolymer microcapillary strips enabled direct non-invasive optical monitoring of size and morphology of microalgal populations, including growth kinetics, single- cell morphology analysis, and cell cycle events from just 10 ^l of medium / sample, matching cell events observed with flow cytometry. This method provides a microanalytical solution for in-situ water management and high-throughput microalgal productivity assessment.

[0033] In an embodiment, modelling of growth of the microalgae in the microcapillary strips followed a Monod-kinetics with light intensity being the limiting factor.

[0034] In an embodiment, the disclosed apparatus can be part of a kit for microalgae and cyanobacteria growth investigations and morphology determination; part of a portable sensor for sampling and prediction of HABs; or part of a portable sensor for sampling and determination of water quality.

[0035] In an embodiment, the suitable nutrients are selected from nitrogen, phosphorus, magnesium sulphate, calcium chloride, manganese chloride, zinc sulphate, sodium molybdate, copper sulphate, and mixtures thereof.

[0036] In an embodiment, the inner surface of the microcapillary channel is further covered with citric acid, sodium EDTA, sodium carbonate and / or boric acid.

[0037] In a further embodiment, the inner surface of the microcapillary channel is further covered with copper sulfate, herbicides, antibiotics, and / or algaecides.

[0038] The present disclosure relates to an apparatus for microalgae and / or cyanobacteria cell culture comprising: a unitary body formed from a hydrophobic material; and two or more microcapillary channels extending along the unitary body; wherein the inner surface of the microcapillary channels is coated with nutrients suitable for microalgae and / or cyanobacteria cell culture; and wherein the hydrophobic material is transparent to visible light.

[0039] In an embodiment, the suitable nutrients are selected from a list comprising nitrogen, phosphorus, magnesium sulphate, calcium chloride, manganese chloride, zinc sulphate, sodium molybdate, copper sulphate, or mixtures thereof.

[0040] In an embodiment, the suitable nutrients are nitrogen and phosphorus.

[0041] In an embodiment, the molar ratio between nitrogen and phosphorus in each microcapillary channel ranges from 10:1 to 40:1, preferably is 10:1.

[0042] In an embodiment, the apparatus is sealable in both ends of the two or more microcapillary channels, preferably by silicone grease, lubricating grease, polytetrafluoroethylene-based grease (Teflon-based grease), hydrogel sealants (such as ethylene glycol-based hydrogels, or chitosan-based hydrogels), medical-grade mineral oil or petrolatum (United States Pharmacopeia grade), fluorinated oils, fluorinated greases, vegetable oil-based lubricants, medical-grade polyurethane adhesives and sealants, or mixtures thereof.

[0043] In an embodiment for better results, the two or more microcapillary channels are sealed, preferably sealed by silicone grease, lubricating grease, polytetrafluoroethylene-based grease, hydrogel sealants, medical-grade mineral oil or petrolatum, fluorinated oils, fluorinated greases, vegetable oil- based lubricants, medical-grade polyurethane adhesives and sealants, or mixtures thereof.

[0044] In an embodiment, the hydrophobic material has a refractive index ranging from 1.26 to 1.40, the refractive index being measured at 20 °C with light of wavelength 589 nm

[0045] In an embodiment, the inner diameter of the at least two microcapillary channels ranges from 100 to 250 µm.

[0046] In an embodiment, the inner diameter of the at least two microcapillary channels ranges from 190 to 220 µm.

[0047] In an embodiment, the length of the at least two microcapillary channels ranges from 0.01 m to 2.00 m, preferably from 0.02 m to 0.50 m, more preferably from 0.03 m to 0.05 m. In another embodiment, the length of the at least two microcapillary channels ranges from 0.01 m to 0.03 m.

[0048] In an embodiment, the apparatus comprises three or more microcapillary channels, wherein the microcapillary channels are parallelly arranged.

[0049] In an embodiment, the apparatus comprises at least five microcapillary channels parallelly arranged, preferably ten microcapillary channels parallelly arranged.

[0050] In an embodiment, the hydrophobic material is a fluoropolymer selected from a list comprising fluorinated ethylene propylene (FEP), tetrafluoroethylene hexafluoropropylene vinylidene fluoride(THV), perfluoroalkoxy (PFA), polytetrafluoroethylene (PTFE), ethylene tetrafluoroethylene (ETFE), poly(chlorotrifluoroethylene) (PCTFE), polydimethylsiloxane (PDMS), and mixtures thereof.

[0051] In an embodiment, the inner surface of the at least two microcapillary channels has a hydrophilic inner surface.

[0052] In an embodiment, the inner surface of the at least two microcapillary channels is coated with polyvinyl alcohol.

[0053] In an embodiment, the apparatus is a microcapillary strip.

[0054] An aspect of the present disclosure relates to a kit comprising the apparatus according to any of the previous embodiments.

[0055] The present disclosure also relates to a sensor comprising the apparatus according to any of the previous embodiments.

[0056] It is also disclosed the use of the disclosed sensor for detection of harmful algal blooms, prediction of harmful algal blooms, or assessment of water quality. In an embodiment, the sensor is used for detection of harmful algal blooms, prediction of harmful algal blooms, or assessment of water quality, in static culture conditions.

[0057] The present disclosure also relates to the use of an apparatus as a reactor for cultivation of microalgae and / or cyanobacteria, preferably in static culture conditions, wherein the apparatus comprises a unitary body formed from a hydrophobic material; and two or more microcapillary channels extending along the unitary body; wherein the hydrophobic material is transparent to visible light.

[0058] In an embodiment, the inner diameter of the at least two microcapillary channels ranges from 100 to 250 µm, and the length of the at least two microcapillary channels ranges from 0.01 m to 2.00 m, preferably from 0.02 m to 0.50 m, more preferably from 0.03 m to 0.05 m. In another embodiment, the length of the at least two microcapillary channels ranges from 0.01 m to 0.03 m.

[0059] In an embodiment, the at least two microcapillary channels are further coated with nutrients suitable for microalgae and / or cyanobacteria cell culture, preferably nitrogen and phosphorus.

[0060] In an embodiment, the molar ratio between nitrogen and phosphorus ranges from 10:1 to 40:1, preferably is 10:1.

[0061] An aspect of the present disclosure relates to a method for culturing microalgae and / or cyanobacteria in an apparatus comprising a unitary body formed from a hydrophobic material and two or more microcapillary channels extending along the unitary body, wherein the hydrophobic material is transparent to visible light, the method comprising the following steps: loading the at least two microcapillary channels with culture medium comprising an inoculum of microalgae and / orcyanobacteria; sealing both ends of the at least two microcapillary channels; placing the apparatus in a growth chamber operating at a suitable temperature and photoperiod; optionally, observing the apparatus under a microscope.

[0062] In an embodiment, the loading step is performed by capillary action.

[0063] In an embodiment, both ends of the at least two microcapillary channels are sealed with silicone grease, lubricating grease, polytetrafluoroethylene-based grease, hydrogel sealants, medical-grade mineral oil or petrolatum, fluorinated oils, fluorinated greases, vegetable oil-based lubricants, medical- grade polyurethane adhesives and sealants, or mixtures thereof; preferably are sealed with lubricating grease.

[0064] In an embodiment, the suitable temperature ranges from 18 to 23 °C.

[0065] In an embodiment, the suitable photoperiod comprises twelve hours of light and twelve hours of darkness, and light intensity of 30 μmol / m2 / s.

[0066] In an embodiment, the at least two microcapillary channels are further coated with nutrients suitable for microalgae and / or cyanobacteria cell culture, preferably nitrogen and phosphorus.BRIEF DESCRIPTION OF THE DRAWINGS

[0067] The following figures provide preferred embodiments for illustrating the disclosure and should not be seen as limiting the scope of invention.

[0068] Figure 1: Schematic representation of an embodiment of cultivation of Parachlorella kessleri. (A) Cultivation in Erlenmeyer flasks, either unsparged or sparged with filtered air; (B) Microphotograph of a microcapillary strip showing top view and cross section of 10 microcapillary channels, each with mean hydraulic diameter of 206 ± 12.6 µm

[0010] (C) Micro-cultivation process of P. kessleri which started by loading cells diluted in medium by capillary action. The colour of the culture drop does not represent the cell concentration used in the experiments. Scale bar 1 mm.

[0069] Figure 2: Embodiment of cell image analysis of the sequential micrographs taken every two days, showing Parachlorella kessleri cells proliferating inside the microcapillary strips. (A) Time-sequence of micrographs taken every 2 days, up to day 10, for the same group of cells; on the right column the red dashed-line rectangles represent the selected section of the microcapillary with cells highlighted in binary image after ImageJ analysis; scale bar represents 100 µm; (B) Distribution of arbitrarily selected P. kessleri cells diameter (µm), from the microphotographs taken over the growth curve time (n=100) (inset). Cell diameter from randomly picked microphotographs (grey) compared with the size distribution obtained from a mixture of cells from different growth stages, with flow cytometry (white);(C) Boxplot representation of all individual cell sizes found in the micrograph selected area at day 2, 4, 6 and 8, + signal refers to isolated values.

[0070] Figure 3: Illustration of results of growth of Parachlorella kessleri cells over 22 days with different cultivation methods.

[0071] Figure 4: Schematic representation of the functionalization of a microcapillary strip with nutrients.

[0072] Figure 5: Embodiment of results of A) Growth curves of Parachlorella kessleri cells in preloaded BG11 medium functionalized microcapillary strips; (1) Functionalized strips with an N:P ratio of 10:1 (grey pentagon); (2) Functionalized strips with N:P ratio of 20:1 (squares); (3) Functionalized strips with N:P ratio of 30:1 (triangle); (4) Functionalized strips with N:P ratio of 40:1 (circle). B) Maximum growth rate achieved in the exponential phase of Parachlorella for 22 days, with the different functionalized strips. C) Growth curves of Parachlorella kessleri cells analysed under different cultivation methods; Cell growth: (1) in Functionalized microcapillary strips with an N:P ratio of 10:1 (corresponding to the pentagon); (2) PVOH coated microcapillary strips; (3) Erlenmeyer flasks in batch (diamonds); (4) Erlenmeyer flasks with mixing by aeration with filtered air (triangles). D) Maximum growth rate achieved in the exponential phase of Parachlorella for 22 days, for the different culturing methods.

[0073] Figure 6: Embodiment of results of A) Growth curves of Parachlorella kessleri cells in microcapillary strips, with different light exposure; Cells growing: (1) under 15 µmol / m2 / s (diamond); (2) without light (circles); (3) in an encapsulated environment, with reduced gas exchanges. B) Maximum growth rate achieved in the exponential phase of Parachlorella for 22 days, with the different light and gas conditions.

[0074] Figure 7: Embodiment of growth of Parachlorella kessleri in an open microcapillary strip (comparative data, no seal) and growth in the apparatus of the present disclosure, i.e., closed pristine microcapillary strip, over a period of three days.

[0075] Figure 8: Embodiment of results of long-term cultivation of A) viable cyanobacteria Oscillatoria sp. filaments cultured in the disclosed apparatus, under controlled suitable conditions of light and temperature, viable after 1 year, with no nutrient renewal (scale bar: 100 µm); B) viable microalga Parachlorella kessleri cultured in the disclosed apparatus under controlled suitable conditions of light and temperature, viable after 2 years, with no nutrient renewal (scale bar: 50 µm).DETAILED DESCRIPTION

[0076] The present disclosure relates to an apparatus for microalgae and / or cyanobacteria cell culture comprising a unitary body formed from a hydrophobic material; and two or more microcapillarychannels extending along the unitary body; wherein the inner surface of the microcapillary channels is coated with nutrients suitable for microalgae and / or cyanobacteria cell culture; and wherein the hydrophobic material is transparent to visible light, and to a kit and sensor thereof. A method for culturing microalgae and / or cyanobacteria in an apparatus comprising a unitary body formed from a hydrophobic material and two or more microcapillary channels extending along the unitary body, wherein the hydrophobic material is transparent to visible light is also disclosed.

[0077] The present disclosure is more particularly described in the following examples that are intended as illustrative only since numerous modifications and variations are possible and will be apparent to those skilled in the art.

[0078] Cultivation and cell concentration determinations in Erlenmeyer flasks

[0079] In an embodiment, it was selected the model organism Chlorella which is a unicellular chlorophyte, with amenable characteristics to enable the exploration of the volumetric and optical characteristics of the microcapillaries. A microalgal strain of Chlorella sp. A2O 1394 was purchased from the company Alga2O. The species name was determined by molecular biology tools. Cells were collected by centrifugation of 0.5 mL culture and disrupted using a mixer mill (MM200 Retsch, Haan, Germany) for 5 min. Genomic DNA was extracted using NucleoSpin Plant II (Macherey-Nagel). The PCR and sequencing service was outsourced to StabVida (stabvida.com). Primers used for obtaining sequences of the 18S rRNA gene included the universal amplification primers used for eukaryotic algae 18S-F and 18S- R and originated a sequence of 1503 bp. Primers used for amplification of rbcL were 1AB_rbcLF and 1AB_rbcLR and originated a sequence of 627 bp. Sequencing reads were assembled with SeqAssem (SequentiX, http: / / www.sequentix.de / software_seqassem.php) and manually edited by visual inspection of sequencing chromatograms. The sequences were loaded into GenBank blastn search engine from NCBI (https: / / blast.ncbi.nlm.nih.gov / ). The combined results from the two gene sequences indicate that the organism is Parachlorella kessleri, a species of microalgae previously known as Chlorella kessleri but with a distinct molecular setup.

[0080] In an embodiment, Parachlorella kessleri was cultured in 250 mL Erlenmeyer flasks with BG11 culture medium (50x freshwater solution, Sigma), preferably for three weeks. Two cultures were set in batch Erlenmeyer flasks, one unsparged and the other sparged with aeration using an air pump providing filtered air (FALC 0.22 µm diameter pore) (Fig. 1A). Each of these cultures was initiated by diluting an inoculum which was pre cultivated in for 3 weeks, with fresh medium to a final cell concentration of 4x105cell / mL. Cultures were placed in a growth chamber (aralab, FC S600PLH) with temperature 18°C, photoperiod 12h:12h and light intensity 30 μmol / m2 / s provided by cool white daylight fluorescent lamps. Aliquots were taken every other day and counted (Brand-Blaubrand Sigma- Aldrich counting chamber) for cell concentration estimations (n=4).

[0081] In another embodiment, a culture of Parachlorella kessleri was left to grow for 3 weeks in 250 mL Erlenmeyer flasks containing BG11 culture medium (x50 freshwater solution, Sigma). Samples from these cultures were then removed and diluted to achieve an approximate starting cell quantity of 4x105cell / mL and loaded onto 3-centimeter long microcapillary strips composed of 10 parallel arrays of microcapillary channels, with 206 ± 12.6 µm mean diameter made of ethylene propylene (Commercially named Teflon®).

[0082] Cultivation of Parachlorella kessleri and optical imaging in microcapillary strips

[0083] In an embodiment, it was used microcapillary strips produced by Lamina Dielectrics Ltd (Billingshurst, West Sussex, UK) consisting of a transparent fluorinated ethylene propylene (Teflon® FEP, Dupont, USA

[0012] ) array of 10 parallel microcapillary channels with 206 ± 12.6 µm mean diameter (Fig. 1B).

[0084] In an embodiment, the strips were used as produced. For this, the loading process required the use of a syringe and a 3D-printed mechanism to facilitate the filling of the microcapillary channels within the microcapillary strip.

[0085] In another embodiment, the inner surface of the microcapillary channels contained within the microcapillary strip were coated with polyvinyl alcohol (PVOH), making the inner surface of the microcapillaries hydrophilic

[0010] , promoting capillarity within them. In another embodiment, PVOH- coated microcapillary film was trimmed into 30 mm long strips. Each cultivation experiment was carried out with at least four replicas. The volume of culture medium inside each 30 mm long strip was 10 µL, corresponding to 1 µL per microcapillary. Aliquots of P. kessleri culture (concentration of 4x105cell / mL) were loaded by capillary action (Fig.1C) and microcapillaries were sealed on both ends using lubricating grease (75003786, Thermo-Scientific™). Nutrients were not renewed in the microcapillary strips during the full extent of the cultivation experiments. Each microcapillary strip was taped to the bottom of a standard plastic Petri dish and observed with an inverted microscope (Zeiss Vert.A1) at regular intervals for quantitation of projected area of the growing microalgal in a pre-defined imaging area. The Petri dish and the microscope stage were setup in order to enable sequential photography in the same location of the strip (Axiocam 208 color, Image software ZEN 3.3 Blue Edition, Zeiss). Micrographs were obtained every other day for 22 days. Once imaged at room temperature (20 °C), strips were returned to the growth chamber to resume exposure to controlled temperature of 20°C, photoperiod of 12h:12h and light intensity of 30 μmol / m2 / s. Sets of strips were incubated in different light conditions: at dark, under low light ~15 μmol / m2 / s and under growth light 30 μmol / m2 / s. An additional set of strips was treated to limit air exchange, by encapsulation with glass slides on top and bottom and grease sealing on the sides.

[0086] In yet another embodiment, the microcapillaries were functionalized by loading with culture medium with different concentrations of nutrients and then lyophilized (Fig. 4). After the loading, thetips of the microcapillary strip are sealed with heat contact and lubricating grease (75003786, Thermo- Scientific™). In an embodiment, four pristine sets of functionalized strips were prepared and preloaded with altered BG11 medium with a varying ratio of nitrogen to phosphorus (N:P). The ratios were 10:1, 20:1, 30:1, and 40:1. The microcapillary strips were afterwards lyophilized. A control set containing strips coated with PVOH was also prepared. All sets were loaded at the same time and allowed to grow for 22 days. Two pristine sets were prepared and incubated under different light conditions: low light with 15 μmol / m2 / s and complete darkness. Another pristine set was prepared with reduced gas exchanges, where the strips were covered with lubricating grease on the sides and placed between two microscope slides.

[0087] In an embodiment, cultivation of microalgae was performed. For each test, a set was prepared, consisting of 4 replicas for each strip type placed onto a petri dish. Sets were then placed in a growth chamber (Aralab, FC S600PLH), with a temperature of 18°C, a photoperiod of 12 hours on and 12 hours off, and light intensity of 30 μmol / m2 / s supplied by cool white daylight fluorescent lamps.

[0088] Determination of cell concentration of Parachlorella kessleri cultivated in microcapillary strips and flow cytometry

[0089] In an embodiment, the mean single cell area and cell concentration inside each microcapillary were determined based on optical microscopy imaging. The mean area of one Parachlorella cell living in the microcapillary channels, Acell, was obtained with ImageJ (NIH, USA) by determining the area occupied by individual cells out of 100 randomly selected cells found in the micrographs, yielding a mean area of 20.20 ± 13.54 µm2for one cell. Cell concentration (cells / mL) was estimated based on image analysis of a pre-set rectangular area, of 88,387.2 µm2, which was analysed for each individual microcapillary and used in all micrographs examined over time (Fig. 2A). The first step consisted in setting a colorimetric threshold in ImageJ that enabled determining the total area occupied by the cells on each microcapillary, teemed Aprojected. The projected area for a 30 mm in length x 206 µm wide microcapillary was estimated as Aprojected = 6,180,000 µm2. Then, determined the total number of cells present in a rectangle of 88,387.2 µm2was determined, at each time point, by dividing the projected area occupied by cells, Aprojected, by the single cell averaged area, Acell. The volume of the culture occupying the rectangle, here called interrogation volume, Vinterrogation, was determined to be 0.01430 µl, given that each 30 mm long microcapillary contained a total of 1 µl. Hence, the cell concentration calculations were performed in all the images obtained along the growth curve as follows (equation 1): (Equation 1)

[0090] In an embodiment, it was performed a measuring validation experiment with cytometric analysis using an Attune NxT (Invitrogen) cytometer. It was determined the cell sizes of a mixed culture consisting of cells from different stages of growth, therefore recreating cell measurements in the microcapillary strips. Forward scatter (FSC) and side scatter (SSC) measurements were taken, and fluorescent gating of algal cells was carried out with the 695 / 40 nm BL3 filter which detects chlorophyll fluorescence. This excludes debris and bleached cells. Cell duplets were excluded by gating side scatter hight SSC-H and side scatter area SSC-A measurements. For determining cell sizes, it was used a mix of polystyrene beads of known sizes (Size Calibration kit F13838, ThermoFisher. To overcome the slight discrepancy of the polystyrene beads refraction index (RI), 1.58-1.8, with that of cells which is estimated to be 1.36-1.46

[0013] , it was used the width parameter (SSC-W), an approach already successfully applied to algal cells

[0013] . Cells were gated in SSC-W and BL3 then mean SSC-W was estimated for the size marker beads of 2 µm, 4 µm, 6 µm, and 10 µm and integrated in the SSC-W profile of P. kessleri for a total of ~10000 analysed cells. The results were compared with cell size estimations performed with image analysis by adjusting the number of bins to 4 and considering that the standard deviation of the bins should not be higher than that of the cytometric analysis.

[0091] Growth modelling of Parachlorella kessleri cells

[0092] The growth of P. kessleri in the microcapillaries and Erlenmeyer followed an exponential growth model for the first 8-10 days of cultivation, described by Equation 2:(Equation 2)

[0093] where N is the cell concentration (cells / mL), µ is the growth rate, and t is time. However, growth rate quickly dropped beyond what would be estimated from a Monod kinetics limited by substrate. Consequently, it was modelled the microalgal growth as a Monod-kinetic limited by light intensity, I (Equation 3): (Equation 3)

[0095] where µ is the growth rate, µmaxis the maximum growth rate, and KIis the light intensity for half of maximum growth rate. As incident light intensity I is kept constant throughout the experiment, it was hypothesised the biggest contribution to decline of µ is the build-up of cell concentration in the medium, which creates a shadow effect to other cells. To captured this the Lambert-Beer law (Equation 4) was considered: (Equation 4)

[0097] where I0is the incident light intensity, N is the cell concentration, ^ is the light extinction coefficient and l is the light path distance. Rearranging Eq. (3) yields:

[0098] (Equation 5)

[0099] where a is an arbitrary constant proportional to the product of ^ and ‘volume weight’ cell concentration, N.

[0100] Combining Equations 1 to 3 with Equation 5, it can be shown the increase in N can be modelled by the following equation: (Equation 6)

[0102] As Equation 6 is non-linear, kinetic parameters KI’and a were calculated for each experimental growth curve by best fitting the model to the experimental using MATLAB R2020b and minimum square differences. In addition, µmaxwas calculated directly from the experimental data by best-fitting data for exponential stage to Equation 2 integrated as follows: (Equation 7)

[0104] where N0 is the initial cell concentration. This assumes the µ = µmax for initial stage of growth, which has been confirmed by experimental.

[0105] The determination of the growth rate enabled the calculation of the divisions per day (div / day), as given by equation 8: (Equation 8)

[0106] In an embodiment, aliquots of P. kessleri culture were loaded in a transparent apparatus by capillary action, as depicted in Fig.1C. Then, microcapillaries were sealed on both ends using lubricating grease, and nutrients were not renewed in the transparent apparatus (microcapillary strips) during the full extent of the cultivation experiments. Each microcapillary strip was taped to the bottom of a standard plastic Petri dish and observed with an inverted microscope at regular intervals for quantitation of projected area of the growing microalgal in a pre-defined imaging area.

[0107] In an embodiment, the portability of the microcapillary strips and the optical transparency of the fluoropolymer made it possible to examine the microcapillary strips with a standard optical microscope and track individual cell dimensions and cell concentration, in real-time. P. kessleri cells were able to grow in the microcapillaries retaining their spherical morphology and green colour throughout the extent of micro-cultivation period, i.e., 22 days, as shown in Fig.2A.

[0108] In an embodiment, the proliferation of cells in the microcapillaries was possible by optically examining the same area of each microcapillary throughout the cultivation time (22 days). Fig.2A shows a time sequence of microphotographs with cells visible after 2, 4, 6, 8 and 10 days inside themicrocapillary channel strip. It was observed an initial group of cells after 2 days in the microcapillary strips (Fig.2A, 2 days) larger in size, in contrast to smaller cells observed on day 10 onwards (Fig.2A, 10 days). Between days 2 and day 4, the cell number stagnated whilst increasing in size, presumably corresponding to a period of acclimatation. Then, between days 4 to 6, some of the large cells were no longer visible, with a dominance of smaller cells. This transition extended until day 10, when no cells larger than 5 ^m could be observed in the population.

[0109] In an embodiment, the mean projected area of a P. kessleri cell cultivated in the microcapillary strips was estimated as 20.189 ± 13.54 µm2, corresponding to a mean cell diameter (n=100) of 6.2 ± 2 µm. P. kessleri cells display a range of diameters from 2.3 µm to 10.1 µm, following a Gaussian Kernel distribution with a Full Width at Half Maximum (FWHM) of 3.56 (Fig. 2B, inset). The cell size range estimations of P. kessleri based on the imaging in microcapillary strips matched well results from a state- of-the-art flow cytometer, with the use of size marker beads (Fig. 2B). The results show that most cells measure 4 to 6 µm and that optic methods (n=100) were able to distinguish cell sizes, which is in the same range as the results found with cytometric analysis (n=10000). Lower resolution was achieved in the smaller cells of 2 µm which were ~4% less detected by optical methods. It was noted that cell diameter was not constant over time but instead it varied according to the cell growth stage. In Fig.2C, it is shown how cell diameters changed as a function of time. Cells start with an average diameter of 6.07 ± 1.73 µm, followed by an increase to 7.82 ± 1.93 µm, which then reached a smaller diameter of 4.67 ± 0.1.69 µm, at day 6, remaining steady at 4.28 ± 0.65 µm, until day 8. These observations were commensurate with previous observation of cell cycle for Chlorella

[0014] , reporting that in the initial days of cultivation, large mother cells are formed, then followed by a stage when mature mother-cells go through an autospore generation.

[0110] In an embodiment, the resolution of the setup enabled to determine which individual cells correspond to actual autospores by following their morphology and size over time. Eight cells were identified as autospores and could be measured until the autospore releasing day. These cells measured 7.13 ± 1.21 µm, at day 2, then enlarged to 8.37 ± 0.76 µm, at day 4, and to 9.01 ± 0.33 µm, at day 6, after which they could not be traced due to cell disruption upon autospore release (Fig.2A).

[0111] In another embodiment, it was compared the growth using microcapillary strips cultivation of P. kessleri cells with batch unsparged and air sparged Erlenmeyer flasks.

[0112] The increase in cell concentration (N) during the cultivation of P. kessleri in batch Erlenmeyer flasks and microcapillary strips were summarised in Table I. As expected, continuous sparging by aeration for 22 days yielded a higher increment in number of cells, promoting cell division and population growth, compared to stagnant cultures (p<0.05). Specifically, sparged cultures yielded 3.1 x107± 4.1 x105cells / mL after 22 days, corresponding to 8-fold increase in cells number since day 0,compared to only a 4-fold increase observed in unsparged culture, with 1.8 x106± 2.1 x105cells / mL in day 22 (Table I). Thus, the growth of microalgae in a unsparged conditions is highly inhibited; injecting gas into the culture system enables the microalgae to grow at maximum rate. This is in line with previous studies that identified mixing as an important factor in microalgal cultivation when large biomass yields are required [1],

[0015] . Indeed, all cultivation and micro-cultivation methods proposed to date include a degree of mixing, ensuring constant access to nutrients and dissolved gases. Surprisingly, cultivation in the microcapillary strips, with no nutrient renewal nor mixing (static microcapillary strips), maximum growth was obtained without the supply of gas nor medium. The achieved a cell concentration was much higher than that of the unsparged cultures (p<0.05) and similar to the cell concentration obtained with the sparged cultures, with nearly 3x107cell / mL after 22 days in culture.

[0113] Figure 3A depicts three stages of growth, in agreement with previous reports [6]: an initial stage of exponential growth from day 0 to day 10, followed by linear growth from day 10 to day 16-18, and finally a declining stage until day 22 (Fig.3A). In particular, Figure 3 represents an embodiment of results of growth of Parachlorella kessleri over 22 days with different cultivation methods. Figure 3A depicts cell growth with different cultivation tests fitted with a modified Monod´s model (dashed lines) (1) Erlenmeyer flasks, unsparged (diamonds) (n=4); (2) Erlenmeyer flasks, sparged with mixing by aeration with filtered air (squares), (n=4); (3) Microcapillary strips in regular growth conditions, under 30 μmol / m2 / s (circles), (n=3); (4) Encapsulated microchannel strips with limited gas exchange (star), (n=6); (5) Microcapillary strips in low light, under 15 μmol / m2 / s (triangles), (n=2); (6) microcapillary strips in darkness (cross), fitting of the model was not possible due to end of growth after an exponential start. Figure 3B shows the rate of increase of N over time, showing the model is able to predict exponential until ^day 10. Figure 3C depicts the growth rate predicted by the model vs experimental data, showing maximum growth rate until N ^ 107cells / ml. For the scope of the present disclosure, “regular growth conditions” refer to a temperature of 18°C, a photoperiod of 12 hours on and 12 hours off, and light intensity of 30 μmol / m2 / s supplied by cool white daylight fluorescent lamps.

[0114] The exponential growth usually occurred at the expense of available nutrients, followed by a linear growth phase and finally a declining growth phase as a consequence of limited growth [4], [6],

[0016] . Interestingly, the evolution of cell concentration in the microstrips at dark did not follow a growth curve. Cells started to divide and a higher cell concentration of 2.09 x106± 3.94 x104was achieved at day 4, likely due to metabolic resources from previously given light conditions (Table I). Then the concentration dropped to 1.63 x106± 4.17 x105cells / mL and remained relatively steady until day 16, after which the concentration started to lower further, due to cell death (Fig.3).

[0115] Table I – Embodiment of cell concentration of P. kessleri cultivated in Erlenmeyer flasks in still (unsparged) and aerated (sparged) by continuous bubbling with filtered air, and in microcapillary strips.Still and aerated cultures were sampled and cells counted every other day with a counting chamber and cell concentration in microcapillary strips were determined by direct optical interrogation and image analysis of individual microcapillaries. Cultivation in microcapillary strips was performed also with gas exchange limitation (encapsulated strips) and in different light conditions, regular light 30 μmol / m2 / s, low light 15 μmol / m2 / s and dark. (a) to (e) denote significantly different (p<0.05), analysis of variance performed with ANOVA in MATLAB R2020b. Culture Erlenmeyer Erlenmeyer Strips Strips Strips low Strips (days) unsparged sparged Encapsulated light dark0 4.8 x105± 3.7 x105± 3.0 4.9 x105± 4.4 x105± 4.4 x105± 4.7 x105± 8.1 x103x1041.6 x1051.8x1041.8x1043.7x1042 5.3 x105± 3.2 x105± 8.8 8.5 x105± 1.4 x106± 5.3 1.2x106± 3.7 1.4 x106± 1.7 x104x1043.1 x105x105x1052.1 x1054 5.8 x105± 1.0 x106± 2.2 1.6 x106± 2.3 x106± 9.3 9.7 x105± 2.1 x106± 2.2 x104x1047.1 x105x1059.4 x1054.8 x1046 9.2 x105± 2.0 x106± 2.6 4.5 x106± 4.2 x106± 1.2 1.2 x106± 1.6 x106± 6.6 x104x1052.6 x106x1068.8 x1055.1 x1058 1.1 x106± 3.6 x106± 2.0 7.7 x106± 6.8 x106± 1.8 2.8 x106± 1.8 x106± 4.6 x104x1053.4 x106x1062.2 x1065.7 x10510 1.1 x106± 9.8 x106± 3.0 1.3 x107± 9.0 x106± 2.4 4.5 x106± 1.6 x106± 4.3 x104(a) x105(b) 4.8 x106(b) x106(b) 3.3 x106(a) 7.9 x105(a) 12 1.2 x106± 1.7 x107± 6.9 1.6 x107± 1.1 x107± 4.8 6.5 x106± 1.7 x106± 8.6 x104x1055.1 x106x1063.6 x1067.5 x10514 1.1 x106± 1.6 x107± 4.0 1.9 x107± 1.3 x107± 2.6 9.3 x106± 1.7 x106± 3.4 x104x1056.9 x106x 1064.7 x1066.0 x10516 1.7 x106± 2.5 x107± 3.7 2.2 x107± 1.5 x107± 3.1 1.3 x107± 1.7 x106± 1.0 x105x1055.8 x106x1065.0 x1067.1 x10518 1.4 x106± 2.5 x107± 8.2 2.3 x107± 1.7 x107± 2.8 1.4 x107± 1.6 x106± 8.7 x104x1046.5 x106x1066.0 x1066.7 x10520 1.7 x106± 2.6 x107± 1.1 2.6 x107± 1.8 x107± 3.4 1,6 x107± 1.5 x106± 4.8 x104x1066.4 x106x1066.9 x1066.1 x10522 1.8 x106± 3.1 x107± 4.1 2.7 x107± 1.9 x107± 3.3 2.0 x107± 1.6 x106± 2.1 x105(c) x105(d) 7.3 x106(d) x106(e) 8.7 x106(e) 5.8 x105(c)

[0116] In an embodiment, the kinetics of growth of microalgae was modelled using a Monod´s adapted function described in equation 6, with growth rates estimated from best fitting of the model to experimental data and summarised in Table II. The maximum growth rates determined from the exponential growth stage of the growth curves with unsparged Erlenmeyer, sparged Erlenmeyer and microcapillary strips were estimated as µmax = 0.12 ± 0.00, 0.35 ± 0.02 and 0.37 ± 0.05 day-1, respectively (Table II), confirming that unsparged culture yielded the lowest growth and also that sparged culture growth is similar to growth in capillary strips, also seen in Table I (p<0.05). These values are in line with other reports with growth-promoting conditions for large scale cultivation of Chlorella [5], [6].

[0117] In an embodiment, the effect of gas permeability of the microstrips was confirmed by following cell growth in gas-limited (encapsulated) conditions (Fig. 3A). At exponential growth, cell numbers in encapsulated microcapillary strips were evolving similarly (p<0.05) to non-encapsulated microstrips,with N=9.0 x106± 2.40 x106cells / mL and 1.3 x107± 4.8 x106cells / mL respectively at day 10 (Table I). Then growth rate started to reduce, with maximum growth rate in encapsulated strips µmax=0.29 day-1, which is lower than µmax=0.37 day-1in microstrips with unlimited gas exchange conditions (Fig. 2B and Table II).

[0118] In another embodiment, the effect of light exposure was evaluated. The microcapillary strips perform well regarding light exposure. The cultures in microcapillary strips exposed to reduced light intensity followed a similar growth curve (Fig. 3A), however cell concentration over time was highly impacted by reduced light, as seen in Table I. After 22 days, the culture in low light strips achieved 2.0 x107± 8.7 x106cells / mL, which was significantly lower (p<0.05), about 33%, than N-obtained in fully exposed strips (Table I), as a consequence of a slower growth rate of µmax=0.25 / day (Table II). Notably, the cell concentration achieved with unsparged experiment was identical (p<0.05) of the dark microstrips, e.g. at day 22 densities were respectively 1.8 x106± 2.1 x105cells / mL and 1.6 x106± 5.8 x105cells / mL (Table I). This indicates that the unsparged cultivation system may possibly display light limitations. Without mixing, the cells proliferate and accumulate at the bottom of the flask and light distribution is therefore not homogeneous, which hampers photosynthetic activity and overall cell function, resulting in impaired growth. Parameter a in Table II, estimated from best-fitting Equation 6 to experimental data, has a quantitative meaning. In general, it was observed a reduction in µ for N ^ 107cells / ml, which multiplied to values of a = 1.00×10-7-1.30×10-7yields a.N =1.00-1.30, which means µ reduction happens when -log10(I / I0) = 1.00-1.30, which mean around 90% drop in incident light. This is usually the criteria used in literature for light-limiting photocatalysis and confirms validity of the used kinetic model.

[0119] The good gas permeability of FEP material, which has shorter diffusion distance for CO2 to reach the cells in microfluidic strips, allowed a good growth of microalgae in the microcapillary strips as compared to the unsparged Erlenmeyers.

[0120] In an embodiment, it was not possible to fit Equation 6 to the data retrieved from the microcapillary strips incubated in the dark because the culture did not follow the same kinetic behaviour, due to light restrictions. The culture achieved N=1.7 x 106cell / mL, statistically not different to the growth observed with the unsparged Erlenmeyer and low light at day 10 (Table I). After day 10, growth ceased with cell concentration reaching a plateau at ~1.7 x106cell / mL, after which N started to decline.

[0121] In an embodiment, the parallel configuration of the microcapillary strips prevents the shadowing. The configuration of the microfluidic strips also enables larger light exposure, provided by the tubular and transparent surface area of the microcapillary strips, since light and CO2supply is a key aspect in biomass growth and in achieving high cell concentrations [6]. The microcapillary growth togrowth in lab scale conditions were benchmarked, rather than larger scale conditions due to the possibility to further develop the device as a long-term device aimed for low maintenance conditions such as a laboratory growth chamber or an outdoor collector / sensor.

[0122] Table II – Growth parameters retrieved from the Monod´s adapted model, slopes of logarithmic data (µmax) determined for the exponential growth phase of all cultivation experiments. Erlenmeyer batch cultures, unsparged or sparged with filtered air, microcapillary strips with regular light 30 μmol / m2 / s, encapsulated microcapillary strips with limited gas exchange, microcapillary strips with low light 15 μmol / m2 / s and microcapillary strips at dark. µmaxis the maximum growth rate; a is an arbitrary constant proportional to the product of ^ (light extinction coefficient) and ‘volume weight’ cell concentration N; KI´ is the light intensity for half of maximum growth rate considering the shadowing effect of cells. Cultivation Method n µmax Mean µmaxMeanKI’Mean KI’Batch unsparged 1 0.12 0.12 ± 0.00 1.00 x10-71.00 x10-7± 0.00 0.90 0.9000 ± 0.0000 2 0.12 1.00 x10-70.90 3 0.12 1.00 x10-70.90 4 0.12 1.00 x10-70.90 Batch sparged 1 0.38 0.35 ± 0.02 1.25 x10-71.25 x10-7± 0.00 0.0085 0.0074 ± 0.0007 2 0.35 1.25 x10-70.0065 3 0.34 1.25 x10-70.0075 4 0.34 1.25 x10-70.0070 Strip 1 0.34 0.37 ± 0.05 1.05 x10-71.00 x10-7± 0.00 0.010 0.0431 ± 0.0293 2 0.43 1.05 x10-70.055 3 0.33 1.05 x10-70.085 Strip encapsulated 1 0.30 0.29 ± 0.06 1.25 x10-71.30 x10-7± 0.21 x10-70.150 0.0709 ± 0.0630 2 0.29 1.25 x10-70.024 3 0.21 1.25 x10-70.025 4 0.39 1.05 x10-70.130 5 0.31 1.25 x10-70.075 6 0.23 1.75 x10-70.0215 Strip low light 1 0.29 0.25 ± 0.05 1.25 x10-71.13 x10-7± 0.13 x10-70.100 0.0565 ± 0.0435 2 0.20 1.00 x10-70.013

[0123] In an embodiment, micro-cultivation of P. kessleri over 22 days in ‘dip stick’ microcapillary strips delivered optimum microalgae growth without nutrient renewal nor physical aeration and an effective exposure to light, with a growth rate comparable to culture in an aerated Erlenmeyer bioreactor.Microalgal growth was highly dependent on light intensity as confirmed by the modified Monod kinetic model. The optical properties of FEP microcapillaries enabled combined accurate determination of cell concentration and size using an optical microscope and image analysis of the projected occupied area. Although P. kessleri cells within the microcapillaries were grown in unmixed conditions, the maximum growth rate observed during exponential stage was found similar to that observed for a fully aerated Erlenmeyer bioreactor, 0.37 day-1and equal to 0.35 day-1respectively. The confined microenvironment of the microcapillaries enabled efficient use of the dissolved CO2by the cells, even when gas exchange was limited. Counting and morphological monitoring of microalgae were benchmarked against state-of- the-art flow cytometry, demonstrating the possibility of simultaneous acquisition of different growth parameters with simple image analysis. The simplicity of manipulation, cultivation, light permeability, and imaging combined with portability and customization possibilities of Teflon FEP microfluidic capillary strips offers an advanced bioanalytical solution for drinking water management and screening of microalgal productivity assessment.

[0124] In a further embodiment, instrument-free micro-cultivation of microalgae in the microcapillary strips, without the need for a lab incubator and optical microscope, makes the disclosed system methodology compelling for in-field settings.

[0125] In another embodiment, the strips were functionalized by pre-loading each of the microcapillaries with different nutrient concentrations, in 4 replicas. Nitrogen and phosphorus (N:P) are the two main macronutrients, required by microalgae to grow and develop, the commercial BG11 medium, and the common recipes used to produce it, have an approximate N:P ratio of 30:1. Cells’ growth and the exponential phase of Parachlorella kessleri could be obtained by optically interrogating the microalgae inside the microcapillary strips.

[0126] In an embodiment, the 4 replicas followed the growth phases mentioned in previous embodiments. I.e., during the first ten days, the growth was exponential, from day 10 to day 18, the growth was linear, and from day 18 to day 22, there was a decline (Figure 5A). These stages are related to nutrient availability, from day 18 there’s a decrease in growth rate caused by nutrient depletion, and in normal conditions after day 22 usually, the cells tend to spend their last resources before they die. High nutrient concentration did not improve cell growth, as the ratio with a higher maximum growth rate (µmax) was the ratio with 10:1 (hexagrams), reaching 0.35 / day (Figure 5B), and a final cell concentration at day 22 of 4.07 x107± 5.59 x106cell / mL, ratio 20:1 (squares) reached a growth rate of 0.32 day-1(Figure 5B) and a final concentration of 2.92x107± 8.85 x106cell / mL. The molar ratios 40:1 (circles) and 30:1 (triangles) reached a maximum growth rate (µmax) of respectively 0.24 and 0.16 day-1(Figure 5B), in day 12 the ratio 30:1 (triangles) strips surpassed the curve of the ratio 40:1 (circles) despite having a lower maximum growth rate in the initial 10 days of the growth curve and reached a final cell concentration at day 22 of 2.35 x107± 1.38 x107cell / mL.

[0127] Figure 5C compares the results obtained in the presence of a N:P ratio of 10:1 and the Erlenmeyer aerated and batch growth curves produced in the previous embodiments. The growth of PVOH microcapillary strips followed a similar trend as described in previous embodiments. The maximum growth rate (µmax) was 0.33 day-1, and the final cell concentration was 3.32x107± 3.55 x106cell / mL. Despite the slight difference in growth rate, the results were consistent with those of the aerated cultures.

[0128] In an embodiment, when using functionalized strips with N:P 10:1, it was achieved a higher final cell concentration of approximately 1x107cell / mL compared to the aerated and PVOH microcapillary strips methods, and approximately 25 times higher when compared to the batch culture.

[0129] In an embodiment, without the PVOH coating the low light strips (diamonds) (Figure 6A), at the end of the linear phase (Day 22) reached a cell concentration of 2.10x107± 3. 42x106cells / mL, almost half of the functionalized microcapillary strips with N:P 10:1, with full light exposure conditions, and with a lower growth rate µmax= 0.28 / day (Figure 6B). The growth rate value mentioned here shows that light has a significant impact on cultures. However, it also suggests that the presence of PVOH can affect how light reaches the cells. This value is almost twice the one obtained in previous embodiments (Figure 3A), where microcapillary strips were used under similar conditions, but with the only difference being the absence of PVOH, thus showing that that the presence of PVOH affects cell growth. In a further embodiment, the dark microcapillary strips showed similar behaviour as the dark strips with PVOH obtained in the previous embodiments (Figure 3A). They had a low growth rate with a µmax of 0.08 day-1(as shown in Figure 6B) and reached a cell concentration of approximately 1.08 x 106cells / mL by day 10. The cell concentration remained relatively stable with small variations until day 22.

[0130] Surprisingly, the microcapillary strips are a much more complete culture method when compared to the common ones, as they provide better results, on a smaller scale, with less complementary equipment, and are cheaper. The small size of the strips also opened the possibility of being used in the field, to collect and grow microalgal cells. However, the use of such strips would be hindered by the lack of nutrients. In the laboratory, it is feasible to control nearly all factors affecting cell growth, whereas in the field, this becomes nearly impossible. This is solved by the functionalization of the strips with suitable nutrients. The ideal nutrient ratio of N:P in the microcapillary strips is between 30:1 – 40:1, with ideal conditions, usually obtained in the laboratory, but with BG11 medium functionalization in the strips, this system can be easily applied in the field, controlling the nutrient levels in the water.

[0131] In a further embodiment, the cultivating utility is accomplished by loading a cell suspension (in cultivation medium) of the desired species, sealing the ends with grease and placing it in regular culture conditions including light, for long-term preservation (superior than 1 year). The strip may be of longerlength, with the possibility of rolling and standing laterally, with several meters occupying a few centimetres of bench, therefore enabling storage of a larger inoculum of the desired species.

[0132] Surprisingly, sealing the ends of the microcapillary strips improves the culture outcome and allows culture for long periods of time. As depicted in Figure 7, Parachlorella kessleri maintain good viability and growth over time when cultured in a sealed apparatus (second row), where the loaded microorganisms remain alive and reproduce well by cell division inside the microcapillary. When microorganisms were cultured in an open system, when no seal exists (top row, comparative data), air enters, and viability of cells is compromised. Additionally, the disclosed apparatus allowed long-term cultivation of different microorganisms (Figure 8).

[0133] The support and funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation programme (grant agreement No.947897).

[0134] For the scope and interpretation of the present disclosure it is defined that “room temperature” should be regarded as a temperature between 15-30 °C, preferably between 18-25 °C, more preferably between 20-22 °C.

[0135] The term "comprising" whenever used in this document is intended to indicate the presence of stated features, integers, steps, components, but not to preclude the presence or addition of one or more other features, integers, steps, components or groups thereof.

[0136] The disclosure should not be seen in any way restricted to the embodiments described and a person with ordinary skill in the art will foresee many possibilities to modifications thereof. The above- described embodiments are combinable.

[0137] The following dependent claims further set out particular embodiments of the disclosure. References: [1] M. I. Khan, J. H. Shin, and J. D. Kim, “The promising future of microalgae: Current status, challenges, and optimization of a sustainable and renewable industry for biofuels, feed, and other products,” Microb. Cell Fact., vol.17, no.1, pp.1–21, 2018, doi: 10.1186 / s12934-018-0879-x. [2] R. Amaral et al., “Ion-driven communication and acclimation strategies in microalgae,” Chem. Eng. J., vol.473, no. June, 2023, doi: 10.1016 / j.cej.2023.144985. [3] P. R. F. Rocha et al., “Collective electrical oscillations of a diatom population induced by dark stress,” Sci. Rep., vol.8, no.1, pp.1–8, Dec.2018, doi: 10.1038 / s41598-018-23928-9. [4] A. Sánchez-Bayo, V. Morales, R. Rodríguez, G. Vicente, and L. F. Bautista, “Cultivation of Microalgae and Cyanobacteria: Effect of Operating Conditions on Growth and Biomass Composition,” Molecules, vol.25, no.12, pp.1–17, 2020, doi: 10.3390 / molecules25122834. [5] P. Bodénès, H. Y. Wang, T. H. Lee, H. Y. Chen, and C. Y. Wang, “Microfluidic techniques for enhancing biofuel and biorefinery industry based on microalgae,” Biotechnol. Biofuels, vol.12, no.1, pp. 1–25, 2019, doi: 10.1186 / s13068-019-1369-z.[6] E. Lee, M. Jalalizadeh, and Q. Zhang, “Growth kinetic models for microalgae cultivation: A review,” Algal Res., vol.12, pp.497–512, 2015, doi: 10.1016 / j.algal.2015.10.004. [7] J. A. Garrido-Cardenas, F. Manzano-Agugliaro, F. G. Acien-Fernandez, and E. Molina-Grima, “Microalgae research worldwide,” Algal Res., vol. 35, no. May, pp. 50–60, 2018, doi: 10.1016 / j.algal.2018.08.005. [8] A. B. Alias et al., “Microfluidic Microalgae System: A Review,” Molecules, vol. 27, no. 6, 2022, doi: 10.3390 / molecules27061910. [9] H. S. Kim, T. P. Devarenne, and A. Han, “Microfluidic systems for microalgal biotechnology: A review,” Algal Res., vol.30, no. July 2017, pp.149–161, 2018, doi: 10.1016 / j.algal.2017.11.020.

[0010] N. M. Reis, J. Pivetal, A. L. Loo-Zazueta, J. M. S. Barros, and A. D. Edwards, “Lab on a stick: Multi- analyte cellular assays in a microfluidic dipstick,” Lab Chip, vol. 16, no. 15, pp. 2891–2899, 2016, doi: 10.1039 / c6lc00332j.

[0011] S. H. Needs et al., “ Moving microcapillary antibiotic susceptibility testing (mcAST) towards the clinic: unravelling kinetics of detection of uropathogenic E. coli , mass-manufacturing and usability for detection of urinary tract infections in human urine ,” Sensors & Diagnostics, vol.2, no.3, pp.736–750, 2023, doi: 10.1039 / d2sd00138a.

[0012] DuPont, “DuPont Teflon FEP Fluoroplastic,” Prop. Bull., pp.1–4, 2013, Accessed: Oct. 09, 2023. [Online]. Available: https: / / docs.rs-online.com / 7e7e / 0900766b8065a8bd.pdf

[0013] C. Cho et al., “Study of Optical Configurations for Multiple Enhancement of Microalgal Biomass Production,” Sci. Rep., vol.9, no.1, pp.1–11, 2019, doi: 10.1038 / s41598-018-38118-w.

[0014] M. Yamamoto, M. Fujishita, A. Hirata, and S. Kawano, “Regeneration and maturation of daughter cell walls in the autospore-forming green alga Chlorella vulgaris (Chlorophyta, Trebouxiophyceae),” J. Plant Res., vol.117, no.4, pp.257–264, 2004, doi: 10.1007 / s10265-004-0154-6.

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Claims

C L A I M S1. An apparatus for microalgae and / or cyanobacteria cell culture comprising: a unitary body formed from a hydrophobic material; and two or more microcapillary channels extending along the unitary body; wherein the inner surface of the microcapillary channels is coated with nutrients suitable for microalgae and / or cyanobacteria cell culture selected from a list comprising nitrogen, phosphorus, magnesium sulphate, calcium chloride, manganese chloride, zinc sulphate, sodium molybdate, copper sulphate, or mixtures thereof; and wherein the hydrophobic material is transparent to visible light.

2. The apparatus according to any of the previous claims wherein the suitable nutrients are nitrogen and phosphorus.

3. The apparatus according to the previous claim wherein the molar ratio between nitrogen and phosphorus in each microcapillary channel ranges from 10:1 to 40:1, preferably is 10:

1.

4. The apparatus according to any of the previous claims wherein the apparatus is sealable in both ends of the two or more microcapillary channels.

5. The apparatus according to any of the previous claims wherein the two or more microcapillary channels are sealed, preferably sealed by silicone grease, lubricating grease, polytetrafluoroethylene-based grease, hydrogel sealants, medical-grade mineral oil or petrolatum, fluorinated oils, fluorinated greases, vegetable oil-based lubricants, medical-grade polyurethane adhesives and sealants, or mixtures thereof.

6. The apparatus according to any of the previous claims wherein the hydrophobic material has a refractive index ranging from 1.26 to 1.40, the refractive index being measured at 20 °C with light of wavelength 589 nm.

7. The apparatus according to any of the previous claims wherein the inner diameter of the at least two microcapillary channels ranges from 100 to 250 µm.

8. The apparatus according to any of the previous claims wherein the inner diameter of the at least two microcapillary channels ranges from 190 to 220 µm.

9. The apparatus according to any of the previous claims wherein the length of the at least two microcapillary channels ranges from 0.01 m to 2.00 m, preferably from 0.02 m to 0.50 m, more preferably from 0.03 m to 0.05 m.

10. The apparatus according to any of the previous claims comprising three or more microcapillary channels, wherein the microcapillary channels are parallelly arranged.

11. The apparatus according to any of the previous claims comprising at least five microcapillary channels parallelly arranged, preferably ten microcapillary channels parallelly arranged.

12. The apparatus according to any of the previous claims wherein the hydrophobic material is a fluoropolymer selected from a list comprising fluorinated ethylene propylene, tetrafluoroethylene hexafluoropropylene vinylidene fluoride, perfluoroalkoxy, polytetrafluoroethylene, ethylene tetrafluoroethylene, poly(chlorotrifluoroethylene), polydimethylsiloxane, and mixtures thereof.

13. The apparatus according to any of the previous claims wherein the inner surface of the at least two microcapillary channels has a hydrophilic inner surface.

14. The apparatus according to any of the previous claims wherein the inner surface of the at least two microcapillary channels is coated with polyvinyl alcohol.

15. The apparatus according to any of the previous claims wherein the apparatus is a microcapillary strip.

16. A kit comprising the apparatus according to any of the previous claims.

17. A sensor comprising the apparatus according to any of the previous claims 1-15.

18. Use of a sensor according to the previous claim for detection of harmful algal blooms, prediction of harmful algal blooms, or assessment of water quality, in static culture conditions.

19. Use of an apparatus as a reactor for cultivation of microalgae and / or cyanobacteria, preferably in static culture conditions, wherein the apparatus comprises a unitary body formed from a hydrophobic material; and two or more microcapillary channels extending along the unitary body; wherein the hydrophobic material is transparent to visible light.

20. Use according to the previous claim wherein the inner diameter of the at least two microcapillary channels ranges from 100 to 250 µm, and the length of the at least two microcapillary channels ranges from 0.01 m to 2.00 m, preferably from 0.02 m to 0.50 m, more preferably from 0.03 m to 0.05 m.

21. Use according to any of the previous claims 19-20 wherein the at least two microcapillary channels are further coated with nutrients suitable for microalgae and / or cyanobacteria cell culture, preferably nitrogen and phosphorus.

22. Use according to the previous claim wherein the molar ratio between nitrogen and phosphorus ranges from 10:1 to 40:1, preferably is 10:

1.

23. A method for culturing microalgae and / or cyanobacteria in an apparatus comprising a unitary body formed from a hydrophobic material and two or more microcapillary channels extending along the unitary body, wherein the hydrophobic material is transparent to visible light, the method comprising the following steps: loading the at least two microcapillary channels with culture medium comprising an inoculum of microalgae and / or cyanobacteria; sealing both ends of the at least two microcapillary channels; placing the apparatus in a growth chamber operating at a suitable temperature and photoperiod; optionally, observing the apparatus under a microscope.

24. The method according to the previous claim wherein the loading step is performed by capillary action.

25. The method according to any of the previous claims 23-24 wherein both ends of the at least two microcapillary channels are sealed with silicone grease, lubricating grease, polytetrafluoroethylene- based grease, hydrogel sealants, medical-grade mineral oil or petrolatum, fluorinated oils, fluorinated greases, vegetable oil-based lubricants, medical-grade polyurethane adhesives and sealants, or mixtures thereof; preferably sealed with lubricating grease.

26. The method according to any of the previous claims 23-25 wherein the suitable temperature ranges from 18 to 23 °C.

27. The method according to any of the previous claims 23-26 wherein the suitable photoperiod comprises twelve hours of light and twelve hours of darkness, and light intensity of 30 μmol / m2 / s.

28. The method according to any of the previous claims 23-27 wherein the at least two microcapillary channels are further coated with nutrients suitable for microalgae and / or cyanobacteria cell culture, preferably nitrogen and phosphorus.

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