Introduction

The commonly known plant basil (Ocimum basilicum) is a perennial herb and belongs to the Lamiacaea family. In addition to its main culinary use, the plant and its essential oils are also used in traditional medicine, cosmetics and pharmacology (Bae et al. 2020; Mehring et al. 2020; Yeşil et al. 2020; Zhan et al. 2020). The secondary metabolites oleanolic and ursolic acid produced by the plant are of particular importance, but also linalool and rosmarinic acid. In order to produce the triterpenes oleanolic and ursolic acid under ecologically and economically better conditions, the plant cell cultivation technique can be used instead of cultivating the whole plant. The cambial merismatic cells (CMC), which can be obtained from the cambium of young plants, have great potential as production cell lines. Compared to dedifferentiated cells (DDC), CMCs show faster growth and higher product formation (Ochoa-Villarreal et al. 2015; Mehring et al. 2020).

Another way to increase the productivity of plant cells is to cultivate them as immobilized systems on different surfaces. These systems can be described as “synthetic biofilms” (Schmeckebier et al. 2022). Cultivation of plant cells as such a biofilm can lead to increased productivity compared to suspension cultures (Leonov et al. 2021). The choice of reactor type is also essential for the cultivation of CMCs, as the cells react very sensitively to shear stress. Therefore, cultivation as a synthetic biofilm could offer a low shear stress option, where at the same time possible increased productivity could bring further success.

In order to be able to form a synthetic biofilm, the adhesion of the cells to a surface is of great importance, as well as the surface material and the surface structure (Bos et al. 1999). To assess the ability of cells to form a synthetic biofilm, it makes sense to determine the adhesion strength of a single cell to the surface (Azeredo et al. 2017). To make this possible in the case of Ocimum basilicum CMCs, a suitable cell separation method must first be used as the cells grow in large clusters. So far, only a few methods have been found in the literature that seem suitable for the separation of plant cell cultures. There are methods to separate cells directly from the plant (Wu et al. 2009) or protoplasts from callus cells (Yamada et al. 1972), but these turned out to be unsuitable for this application. The difficulty of cell separation of plant cell cultures lies in dissolving the agglomerated clusters without destroying the cells themselves. Therefore an enzyme based method for separation was developed in which vital single cells are still present at the end of the procedure. After cell separation, the singulated cells can then be used for various purposes, such as determining a single cell adhesion measurement, as is also carried out in this short communication. In addition to this application, the single cells can also be used for submerged cultivation, as this improves nutrient accessibility, or for microscopic investigations, such as the staining and analysis of cell organelles.

Materials and methods

Strain maintenance and pre-culture

The plant-derived cambial meristematic culture from Ocimum basilicum (Mehring et al. 2020) was maintained on agar plates with LS medium (30 g l-1 sucrose, 4.4 g l-1 LS-Media (Linsmaier and Skoog 1965), 1 g l-1 MES buffer, 1 g l-1 2,4-dichlorphenoxyacetic acid, pH adjusted to 5.7 with sodium hydroxid), with 5.55 g l-1 plant agar, which were renewed every 4 weeks. These cell cultures were stored in an incubator (Binder FP 115, Fa. BINDER GmbH, Tuttlingen, Germany) at 29 °C in the dark. For the pre-cultures, approx. 4 g l-1 of the wet biomass was transferred to a 300 ml Erlenmeyer flask without baffles with 50 ml LS medium. Incubation followed in a shaking incubator (Multitron, Infors HAT, Bottmingen, Switzerland) at 28 °C, 120 rpm in the dark for 14 days. After incubation for 14 days, approx. 15 g l-1 wet biomass was present.

Cell separation

For cell separation, the pre-culture was filtered through a polypropylene filter (Spectra/Mesh, Cole-Parmer, United Kingdom) with a pore diameter of 150 μm. The retentate was incubated for 1.5 h in 10 ml enzyme solution (0.025% (m v− 1) Macerozym R-10 (Duchefa Biochemie, Netherlands) in LS medium) in an overhead shaker (Intelli-Mixer RM-2 L, LTF, Germany) and then filtered again with the same type of polypropylene filter. The filtrate of the second filtration was washed several times with LS medium and then added to the filtrate of the first filtration. For the washing steps, cells were separated by centrifugation (Rotina 48R, Hermie AG Hettich, Gosheim, Deutschland) at 4000 rpm for 4 min. The cells obtained by the filtration steps in the permeate were resuspended in 50 ml LS medium in 300 ml conical flasks and incubated in a shaking incubator (Multitron, Infors HAT, Bottmingen Schweiz) at 29 °C, 120 rpm for 7 days in the dark. Then the cell suspension was filtered again with the polypropylene filter and the filtrate was centrifuged with a sucrose gradient (specified concentrations, listed from bottom to top, 500 g l− 1, 400 g l− 1, 200 g l− 1 and 0.1 M NaCl). The separated cell fraction (lowest layer) was resuspended in 50 ml LS medium.

Fluorescence staining

Fluorescence microscopy images were taken to quickly check the viability of the cells and the degree of separation. To acquire fluorescence microscopic images, the plant cells were stained with the fluorescent dyes fluorescein diacetate (FDA) (excitation wavelength 488 nm, emission spectrum 500–520 nm) and calcofluor white (Calc) (excitation wavelength 405 nm, emission spectrum 425–475 nm). 0.2 ml of the cell suspension were incubated with 0.3 ml of 10% [v v− 1] Calcoflour solution in Sörensen buffer pH 8 and with 0.1 ml of % [v v− 1] FDA (0.22 mg ml− 1 in acetone) with the addition of 0.6 ml of LS medium for 10 min in the dark. To prevent further reactions of the FDA, the samples were put on ice. The fluorescence images were then taken under the microscope (Eclipse Ni, Nikon, Minato, Japan). FDA is converted from viable cells to fluorescein, so that viable cells are stained green, while Calc attaches to the cell wall and thus stains all cells blue.

Resazurin assay

To evaluate cell vitality after cell singulation, Resazurin assay was performed according to Mehring et al. (2021). The biomass before cell singulation, immediately after cell singulation and 7 days after cell singulation was weighed (approx. 100 mg) and placed in the wells of a 96-well microtitre plate containing 180 µl phosphate buffer (pH 7.2). In addition, 20 µl of a 0.2 mg ml− 1 resazurin solution in deionised water was added. Wells containing only resazurin were also measured as a control. Measurements were then made continuously at 540/560 nm excitation and 590 nm emission in a multilabel plate reader (VICTOR multilabel plate reader, PerkinElmer LAS GmbH, Rodgau, Germany).

AFM single cell measurement

The single cell adhesion measurement was carried out with a Nanowizard III (JPK BioAFM– Bruker Nano GmbH, Germany) with a FluidFM add-on (Cytosurge, Switzerland). The AFM was positioned under an acoustic hood (JPK BioAFM– Bruker Nano GmbH, Germany) and mounted on an active vibration isolation system (Halcyonics-i4, Accurion GmbH, Germany) to minimize the effects of environmental vibrations. A glass sample (microscope glass, Thermo Scientific, soda-lime glass) was used as substrate. Additionally, the Nanowizard III is mounted on an inverted microscope (AxioObserver A1, Zeiss, Germany) to observe the cells and to address the plant cells to be approached with the cantilever. A droplet of diluted plant cell solution was placed on a microscope glass. For SCFS (single cell force spectroscopy), a FluidFM micropipette (Cytosurge, Switzerland) with an aperture of 8 μm and a spring constant of 2 N m− 1 was filled with glycerol, calibrated by the contact-based thermal noise method (Butt and Jaschke 1995; Hutter and Bechhoefer 1993) and brought in close proximity to the glass surface. By applying a pressure of -800 mbar compared to the environment, a single cell was drawn to the aperture of the cantilever. For subsequent measurements, the negative pressure was reduced to smaller values (-200 mbar).

All force-distance curves were recorded in a cell culture medium with a setpoint of 10 nN, a z-speed of 0.5 μm s− 1, and a dwell time of 20 s. Two individual cells were measured, with a total of 100 force-distance curves per contact time.

Data processing

Data processing for the resazurin assay was carried out as described by Mehring et al. (2021). To calculate the emission at 590 nm (\( {\epsilon }_{cal})\), the following formula was used:

$$ {\epsilon }_{cal}=\frac{({\epsilon }_{rec}-{\epsilon }_{ctrl})}{\left(\frac{{\epsilon }_{sat}}{{M}_{res}}\right)}$$

Here, \( {\epsilon }_{rec}\) corresponds to the measured value at 540 nm (top) normalised to the weighed-in bio-moisture mass and \( {\epsilon }_{ctrl}\) to the corresponding measured control emission. \( {\epsilon }_{sat}\) is the steady-state phase of the measurement in which saturation was reached and \( {M}_{res}\) is the molar mass of the resazurin used. The vitality before cell singulation was set to 100% and the corresponding vitality immediately after cell singulation and 7 days after cell singulation were set in relation to this.

The given adhesion force was determined by the largest force relative to the baseline by using the software Mountains SPIP (Digitalsurf, France). All given data points represent the mean value of several measured values at the same conditions. The error bars represent the standard deviation of the values used for the mean values.

Results and discussion

Cell separation and cell vitality

In order to develop the cell singulation method, various possibilities described in the literature were tested, which, however, only resulted in the complete destruction of the cells (QU und Guan Yaqin 2017; Naill and Roberts 2004). The cell separation method now described is divided into some steps, (I) Filtration, (II) Enzyme treatment with washing, (III) Incubation and (IV) Filtration with Density centrifugation (Fig. 1). Since the plant cells in the pre-culture grow to large cell agglomerates (Fig. 2, A), the first step is filtration with a polypropylene filter (I). This separates individual cells and smaller agglomerates from the larger cell agglomerates since the plant cells are on the size scale of about 50–150 μm (Mehring et al. 2020) and a pore diameter of 150 μm is used. In this way, the already existing single cells are not stressed during incubation with the enzyme solution and stay vital (Fig. 2B.1, 2). To evaluate the vitality of the cells, in addition to staining with FDA and Calcoflour white and subsequent acquisition of fluorescence microscopy images, which is useful for rapid vitality verification (Fig. 2B, C.2), vitality was assessed with a resazurin assay (Fig. 3). The resazurin reduction rate is a good measurement of cell vitality, as there is a stable linear relationship between reaction rate and biomass quantity, so that the method used allows a good assessment of cell vitality (Mehring et al. 2021). After filtration of the cells, the cells remaining in the retentate are treated with the enzyme solution (II). The enzyme solution consists of macerozyme, which is also used for the isolation of protoplasts. Macerozyme breaks down pectin and other small cell wall components (Yamada et al. 1972). After this enzyme treatment, the vitality of the cells is reduced to approx. 25% (Fig. 3). This makes the subsequent washing steps to completely separate the enzyme solution even more impotant to avoid further damage to the vitality of the cells. This proves also the introduction of the previous filtration step to separate the vital cells in order not to harm them further. As it was shown that the vitality of the cells is strongly stressed by the enzyme treatment, a new incubation of the cells in the growth medium is added (III). This incubation for 7 days allows the cells to recover so that the vitality of the cells after this incubation has increased again to approx. 71% of the pre-cell singulation value (Fig. 3). After this incubation step, however, cell agglomerates may haveformed again, so that another filtration step (IV) is carried out with a polypropylene filter in order to separate these newly formed agglomerates again and obtain vital single cells (Fig. 2C.1, 2). Although, since the solution now obtained still contains cell debris that could possibly impair subsequent AFM measurements, these must be separated by a final centrifugation step. Here, centrifugation to separate the cell fragments is carried out with the aid of a sucrose gradient. After separation of the cell debris, vital single cells are obtained, which can be used for further measurements (Fig. 2C.2).

Fig. 1
figure 1

Flow diagram of the steps for cell separation. (I) Filtration with a polypropylene filter with a pore diameter of 150 μm. (II) Enzyme treatment (Macerozyme R-10 0.025% (m v-1)) of the cells remaining in the retentate, followed by a washing step (LS medium and centrifugation at 4000 rpm for 4 min). (III) Combining the permeate from step (I) with the cells after step (II) for incubation for 7 days (29 °C, 120 rpm). (IV) Filtration with a polypropylene filter with a pore diameter of 150 μm with subsequent density centrifugation of the permeate (sucrose gradient, 500 g l-1, 400 g l-1, 200 g l-1, 0.1 M NaCl)

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Fig. 2
figure 2

A: microscopic picture of cell agglomerates before cell separation; B: after filtration with polypropylene filter; C: singularized cells after all singulation steps; 1: microscopic pictures, 2: fluorescent picture, stained with FDA and Calcoflour white

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Fig. 3
figure 3

Plot of the percentage of vital cells in relation to vital cells before cell separation, directly after incubation with enzyme solution and after the recovery phase of 7 days after the cell singulation evaluated via resazurin assay, mean value and standard deviation are shown, n = 8

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Adhesion measurement

The adhesion force of a singulated cell was determined with the FluidFM over a contact time of 20 s on a glass sample with approx. 4 nN ± 1.2 nN. In comparison, approximately equal adhesion forces are achieved in the literature using the example of Lactococcus lactis subsp. Lactis under comparable measurement conditions (Hofherr et al. 2020). However, it must be taken into account that the bacterium Lactococcus lactis subsp. Lactis with approx. 0.8–1.4 μm (Hansen et al. 2016) is significantly smaller than the plant cells Ocimum basilicum CMC with approx. 50–150 μm (Mehring et al. 2020). Despite a 100-fold higher cell size, no significantly higher adhesion forces are achieved, so that the adhesion of Ocimum basilicum CMC can be classified as comparatively low. However, the adhesion strength of the plant cells could only be compared with microorganisms, as there are no comparable single-cell adhesion measurements with plant cells on surfaces in the literature so far. In future, the measurement of different plant cell lines will also be of interest, as will the investigation of these on different surfaces.

Conclusion

The described method for cell singulation can be successfully applied to obtain vital single cells of the Ocimum basilicum CMC type. A subsequent adhesion measurement on glass was also possible to determine the adhesion forces of the single cell. Since the literature so far provides only few data on separation methods of plant cell cultures, the cell separation method developed here in combination with the resazurin assay offers a good possibility to transfer this method to other plant cell lines. This will also allow further diverse applications to be explored and further investigation methods to be made possible, such as single cell adhesion measurement.