ORIGINAL RESEARCH
published: 04 July 2022
doi: 10.3389/fsufs.2022.934552
Frontiers in Sustainable Food Systems | www.frontiersin.org 1July 2022 | Volume 6 | Article 934552
Edited by:
Lutz Grossmann,
University of Massachusetts Amherst,
United States
Reviewed by:
Daniele Carullo,
University of Milan, Italy
Michael Sandmann,
Neubrandenburg University of Applied
Sciences, Germany
*Correspondence:
Justus Knappert
Specialty section:
This article was submitted to
Sustainable Food Processing,
a section of the journal
Frontiers in Sustainable Food Systems
Received: 02 May 2022
Accepted: 31 May 2022
Published: 04 July 2022
Citation:
Knappert J, Nolte J, Friese N, Yang Y,
Lindenberger C, Rauh C and
McHardy C (2022) Decay of
Trichomes of Arthrospira platensis
After Permeabilization Through Pulsed
Electric Fields (PEFs) Causes the
Release of Phycocyanin.
Front. Sustain. Food Syst. 6:934552.
doi: 10.3389/fsufs.2022.934552
Decay of Trichomes of Arthrospira
platensis After Permeabilization
Through Pulsed Electric Fields (PEFs)
Causes the Release of Phycocyanin
Justus Knappert1*, Jonas Nolte 1, Natalya Friese 1, Ye Yang1, Christoph Lindenberger 2,
Cornelia Rauh1and Christopher McHardy1
1Department of Food Biotechnology and Food Process Engineering, Technische Universität Berlin, Berlin, Germany,
2Department of Mechanical Engineering/Environmental Technology, Ostbayerische Technische Hochschule (OTH)
Amberg-Weiden, Amberg, Germany
The cyanobacterium Arthrospira platensis is a promising source of edible proteins
and other highly valuable substances such as the blue pigment-protein complex
phycocyanin. Pulsed electric field (PEF) technology has recently been studied as a
way of permeabilizing the cell membrane, thereby enhancing the mass transfer of
water-soluble cell metabolites. Unfortunately, the question of the release mechanism
is not sufficiently clarified in published literature. In this study, the degree of cell
permeabilization (cell disintegration index) was directly measured by means of a new
method using fluorescent dye propidium iodide (PI). The method allows for conclusions
to be drawn about the effects of treatment time, electric field strength, and treatment
temperature. Using a self-developed algorithm for image segmentation, disintegration of
trichomes was observed over a period of 3 h. This revealed a direct correlation between
cell disintegration index and decay of trichomes. This decay, in turn, could be brought
into a direct temporal relationship with the release of phycocyanin. For the first time, this
study reveals the relationship between permeabilization and the kinetics of particle decay
and phycocyanin extraction, thus contributing to a deeper understanding of the release
of cell metabolites in response to PEF. The results will facilitate the design of downstream
processes to produce sustainable products from Arthrospira platensis.
Keywords: Arthrospira platensis (Spirulina platensis), pulsed electric field (PEF), cell disintegration, phycocyanin
extraction, image segmentation, trichome decay, propidium iodide (PI) staining
INTRODUCTION
Due to climate change and the growing world population, which is projected to reach up to 10
billion people by 2050 (Abel et al., 2016), the food industry is confronted with huge challenges.
Therefore, sustainable, high-quality protein sources are needed. Because the production of animal
proteins greatly impacts the environment, vegetable proteins should be preferred for consumption
(Nijdam et al., 2012; Poore and Nemecek, 2018). Besides conventional plant-based protein sources
such as legumes, phototrophic microorganisms (i.e., microalgae and cyanobacteria) and other
valuable cell metabolites have been investigated as a source of proteins (Becker, 2007; Grossmann
et al., 2020). Compared to arable crops, culturing microalgae leads to an up to 10-fold biomass
yield per area while the biomass is characterized by a similar or even higher protein content
Knappert et al. Phycocyanin Release PEF Arthrospira platensis
(van Krimpen et al., 2013). Furthermore, microalgae can be
cultivated on non-arable land or even on water surfaces (Kim
et al., 2016). Among the many species available, Arthrospira
platensis (Spirulina platensis) is a promising candidate as a
novel protein source. This filamentous cyanobacterium is often
mistakenly assigned to eucaryotic microalgae. One characteristic
of A. platensis is the arrangement of cells in a helicoidal,
cylindrical trichome. The cells are divided by cross cell walls,
and their division leads to the elongation of the trichome
(Tomaselli, 1997). The proliferation of trichomes is triggered by
their breakage. Arthrospira can accumulate proteins by up to 70%
of its biomass (Oliveira et al., 1999). In addition, protein from A.
platensis is advantageous for human nutrition, since it contains all
the essential amino acids (Becker, 2013). Recent studies revealed
good techno-functional properties of cell proteins i.e., good
emulsifying, foaming, and gel-forming capacity (Benelhadj et al.,
2016; Bertsch et al., 2021; Böcker et al., 2021:Ramírez-Rodrigues
et al., 2021).
One particularly valuable protein from A. platensis is
the blue-pigment protein phycocyanin (PC). It belongs to
the class of phycobiliproteins and acts as an accessory
pigment in photosynthesis. Together with other phycobilins,
allophycocyanin (APC), and phycoerythrin (PE), PC is located
in phycobilisomes at the surface of the thylakoid membrane
(Stadnichuk et al., 2015). It forms the major fraction of phycobilin
proteins and can account for up to 20% of the biomass of A.
platensis (Christaki et al., 2015). Several properties make PC a
valuable molecule when extracted, increasing the overall value of
the biomass. Recently, PC has received a lot of attention from
the food industry, because it is one of the few natural blue food
colorants (Sigurdson et al., 2017). PC shows thermal instability
at temperatures >50◦C (Böcker et al., 2020), which restricts its
use in foods. Nevertheless, in a high-sugar environment, the
pigment has good thermal stability and is therefore particularly
suitable for use in soft drinks and confectioneries (Martelli et al.,
2014). In addition, various studies suggest that PC has anti-
inflammatory, antiviral, and anti-cancer properties and that it
can provide protection against diabetic nephropathy, making it
interesting to the pharmaceutical industry (Shanab et al., 2012;
Pleonsil et al., 2013; Zheng et al., 2013).
Despite great scientific and public interest in microalgae and
cyanobacteria, the major commercial breakthrough has so far
failed to materialize. This can be attributed to high production
costs, particularly concerning cultivation and downstream
processing. On the other hand, the market for A. platensis is
relatively small, and it is mostly offered in the form of powder
or tablets as a dietary supplement. One approach to overcoming
this bottleneck is the implementation of biorefinery concepts.
Similar to an oil refinery, the aim here is to process as many
cell ingredients as possible into different products (Chew et al.,
2017; Caporgno and Mathys, 2018; Carvalho et al., 2020). In the
case of A. platensis, i.e., PC could be extracted and purified while
the remaining cell protein could be offered at a reduced price
for human consumption. For microalgae in general, it has been
shown that this approach can raise the market price for a kilo of
biomass from e.3 to around e30.5 (Ruiz et al., 2016). One crucial
step for this approach is mild cell disruption, which increases
the mass transfer of valuable cell metabolites across the cell
membrane in order to separate them from the residual biomass.
A promising technology for the enhancement of mass transfer
is the technology of pulsed electric fields (PEFs). This technology
is based on the repeated application of high-energy electric fields
(kV cm−1) that are active for a very short time (µs or ns). During
the pulse, spontaneously formed hydrophobic pores, grow and,
above a certain pore size, become hydrophilic and thus permeable
for various cellular substances (Saulis, 2010). A PEF has been
recently investigated in the context of biorefinery approaches
for different microalgae species (Buchmann and Mathys, 2019;
Leonhardt et al., 2020; Papachristou et al., 2021; Carullo et al.,
2022). Compared to other cell disruption methods such as
bead milling, it was claimed that a PEF enables selective cell
permeabilization for water-soluble substances while preserving
cell structure (Goettel et al., 2013; Postma et al., 2016; Lam et al.,
2017b). This in turn should enable facilitated separation of the
cell debris after extraction of valuable substances of interest.
Furthermore, the literature indicates that a PEF seems to be
an energetically more efficient method for cell disruption in
comparison to bead milling or ultrasound, as it enables the release
of water-soluble metabolites at a lower or comparable specific
energy input (Boer et al., 2012; Goettel et al., 2013; Postma
et al., 2017; McHardy et al., 2021). However, in the context of
process design, PEF technology requires a treatment medium
with suitable electrical conductivity, which must fit the geometry
of the treatment chamber used. on the one hand, high electrical
conductivity leads to shorter pulse duration but to increased
electric current and therefore to intensified heating during
treatment on the other hand (Heinz et al., 2001). This could
damage thermally sensitive valuables. Since standard culture
media for A. platensis have an electrical conductivity of around
30 mS cm−1(Aouir et al., 2015), direct treatment in these media
is not possible. The aqueous phase must therefore be replaced,
which can be expected to reduce economic efficiency on an
industrial scale.
Concerning the treatment of A. platensis with PEF, several
studies have shown that the technology is, in principle, suitable
for permeabilizing the cell, thereby releasing compounds such
as PC without extraction of water-insoluble components such
as chlorophyll (Martínez et al., 2017; Jaeschke et al., 2019;
Akaberi et al., 2020; Carullo et al., 2020; Käferböck et al., 2020).
Even though flow cytometry and propidium iodide (PI) staining
have been previously applied to detect the permeabilization of
microalgae (Luengo et al., 2014; Knappert et al., 2020), they
are not applicable to A. platensis because of the large size of
the trichomes. Accordingly, the effect of PEFs on Arthrospira
platensis was always determined indirectly in the cited studies
by measuring the extraction kinetics for different molecules, i.e.,
PC, water-soluble proteins, or carbohydrates. A drawback of this
approach to characterizing the impact of PEFs on cells is the
time-consuming extraction process, which reduces the possible
parameter sets to be investigated. Furthermore, conditions
during the extraction process (e.g., temperature) have a direct
impact on the extraction of the targeted molecule (Akaberi et al.,
2020). This may bias statements regarding treatment efficiency
and make it difficult to determine the true influence of PEFs
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Knappert et al. Phycocyanin Release PEF Arthrospira platensis
on cells. However, knowledge of cell disintegration efficiency is
essential to design processes that ensure complete cell disruption
while protecting the desired product from damage.
The mechanism underlying the release of the aforementioned
molecules is also not conclusively clarified. For example, the
cited studies differ greatly regarding the trend of PC extraction
curves. While Martínez et al. reported that the extraction is
preceded by a lag time of 150 min, Käferböck et al. (2020)
reported the beginning of the extraction process right after PEF
treatment. Jaeschke et al., on the other hand, observed a lag
time depending on specific energy input. The studies contain
contradictory statements regarding whether the preservation
of cell morphology, observed for other microalgae, can be
transferred to A. platensis. Jaeschke et al. reported in their
study the decay of the trichomes over the extraction time for
higher energy inputs (112 and 56 kJ l−1) and only minor
defects for the lowest energy input (28 kJ l−1). In contrast,
Martínez et al., Käferböck et al., and Carullo et al. stated that
trichomes are predominantly preserved. However, none of these
authors indicated the time after PEF treatment at which the size
measurement occurred. A temporally resolved measurement is
missing in all the four studies.
Based on the presented state of research, we hypothesize
that the PEF treatment initially has no influence on the
size of the particles. However, subsequent extraction over a
period of several hours can lead to the disintegration of the
trichomes, which in turn leads to the release of water-soluble
cell metabolites such as PC. If this hypothesis was correct,
the decay of trichomes would depend on treatment intensity,
which could explain the different trends of the extraction curves
presented in the literature. However, to separate the processes
of cell permeabilization, trichome breakdown, and extraction,
direct and uncoupled measures of all phenomena would be
highly beneficial.
In order to test the hypothesis, we first developed a
novel method to directly detect membrane permeabilization
by staining A. platensis with the fluorescent dye propidium
iodide (PI) and fluorescence spectrometry. With this method,
the impact of PEFs on cells of A. platensis can be directly
measured for the first time. This allows for an unbiased
investigation of the effect of PEFs on cells without being affected
by conditions during the extraction process, thus enabling
a better understanding of parameter effects. Accordingly, we
used the novel method to determine the effects of treatment
time, electric field strength, and treatment temperature on
the permeabilization of the cell membrane in the first series
of experiments. The second part of this study addresses the
question of whether PEFs trigger the decay of trichomes.
Therefore, the particle size of trichomes or their fragments
was monitored over a period of 3 h by light microscopy and
digital image processing. Finally, we investigated whether the
possible disintegration of trichomes is correlated temporally with
the release of PC. The aim of this study is, thus, to gain a
better understanding of the relationship between PEF process
parameters and the mechanism for release of water-soluble
contents from cells.
TABLE 1 | Medium composition used in this study for the SOT medium with
original composition and reduced composition.
Solution Substance Original
composition
[g/L]
Reduced
composition
[g/L]
1 NaHCO316.8 0.84
K2HPO40.5 0.5
Distilled water 500 500
2 NaNO32.5 1.25
K2SO41.0 0.5
NaCl 1.0 0.5
MgSO4×7H2O 0.2 0.2
CaCl2×2 H2O 0.04 0.04
FeSO4* 7 H2O 0.01 0.01
Na2EDTA * 2 H2O 0.08 0.08
A5 1.0 1.0
Distilled water 500 500
Trace metal mix A5 H3BO30.286 0.286
MnSO4* H2O 0.15 0.15
CuSO4* 5 H2O 0.222 0.222
Na2MoO4* 2 H2O 0.0079 0.0079
Distilled water 100 ml 100 ml
MATERIALS AND METHODS
Arthrospira Platensis Cultivation
The cyanobacterium Arthrospira platensis (Spirulina), strain
NIES-39, was used in this study. The strain was kept in a
20-ml SOT medium (cf. Table 1) at room temperature and
under constant illumination using fluorescent tubes with a light
intensity of 20 µmol m−2s−1while being placed on a rotary
shaker with a constant rotation frequency of 100 rpm. Light
intensity was measured on the level of the medium surface inside
the flask using a spherical micro quantum sensor and a light
measuring device (ULM-500; Walz, Germany). Every 2 weeks,
1 ml of the culture was inoculated to a fresh medium.
To produce biomass for the experiments, A. platensis was
cultivated in a bubble column photobioreactor with a culture
volume of 1.4 l (reactor volume 1.65 l) and an inner diameter
of 7 cm. Inline measurements of temperature (Biostream
International, Netherlands) and pH (Hamilton, United States)
were conducted using sensor probes, which were connected
to a control unit with an integrated data logger (Biostream
International, Netherlands).
The cultivation occurred in the reduced SOT medium. The
composition of the reduced medium is shown in Table 1
alongside the original SOT composition. The reduced medium
was chosen in order to match the required electrical conductivity
for the PEF treatment and reduce the required dilution factor
after harvesting and concentrating the cells. Previous tests
showed that cell growth in the reduced medium in terms of dry
biomass was comparable to the one in the non-reduced medium
(data not shown). For mixing, the bubble column was gassed with
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Knappert et al. Phycocyanin Release PEF Arthrospira platensis
air at a volume flow rate of 800 ml min−1via a ring sparger.
Because of the reduced amount of carbonate in the medium, its
buffering capacity for the pH is limited. Therefore, the pH was
automatically maintained by enriching the air with CO2using a
CO2mass flow controller (Brooks Instrument, Germany) as soon
as it rose above 9.5. To maintain the temperature at 30◦C, the
bubble column was placed in a water bath, which was tempered
using an external heater. Continuous artificial white light was
applied from two sides with LED panels (LED-Mg, Germany)
with a total area of 0.171 m2. Light intensity was set to 45 µmol
m−2s−1measured at the surface of the bubble column with a
light meter (ULM-500, Walz, Germany).
To determine the cell density of the culture, a correlation
between biomass dry weight and optical density at 750 nm
(OD750) was established. The optical density of several diluted
cell samples was measured in 1-cm cuvettes using a UV/VIS
spectrometer (Lambda 25; Perkin Elmer). The cell dry weight of
the same samples was determined by filtering them through pre-
dried glass microfiber filters (Whatman GF/F, pore size 0.7 µm).
The filters were washed two times with distilled water to remove
salts and then dried for 24 h at 80◦C. For each data point, cell dry
weight and extinction were measured in triplicate. The resulting
correlation was used to calculate the dry biomass of the sample.
The cells were harvested semi-continuously in the late
exponential growth phase. To ensure this, specific growth rate
was determined in a preliminary batch run (data not shown), and
biomass concentration was determined ow times a day over the
entire duration of the experiment. After harvesting, the culture
was diluted with an autoclaved reduced SOT medium such that
the biomass concentration had increased again to the same level
the next day.
PEF Treatment
Sample Preparation
For sample preparation, the dry weight of the harvested cell
culture needed to be adjusted to the intended concentration.
This was conducted in two steps. First, the cells were separated
from the growth medium and concentrated to a higher
concentration than finally needed. Second, the concentrated cells
were diluted and resuspended in a fresh SOT medium, which
was previously adjusted to the desired electrical conductivity
(2 mS cm−1) by adding distilled water (hereafter termed as
resuspension medium).
For this procedure, the dry weight biomass concentration of
the culture was determined by spectrophotometry as described
in Section PEF Treatment. The amount of cell culture needed
for a certain amount of sample was calculated based on mass
conservation; thus, C1V1=C2V2, where C1is the dry biomass
concentration [g l−1] measured inside the culture and V1is the
volume that must be harvested, C2is the desired dry biomass
concentration (factor 1.6 higher concentration than finally
needed), and V2is the desired sample volume. The cell culture
was then filtered through microfilters (pluriStrainer 1 µm;
pluriSelect Life Science, Germany) to separate the cells from
the culture medium. The remaining pellet was rinsed with the
same amount of distilled water to remove the salt. The pellet was
transferred to a 50-ml Greiner tube by washing it off the filter with
a fresh resuspension medium and filling the tube afterward up
to 20 ml. The dry biomass concentration was again determined,
and the required volume of the resuspension medium to reach
the final concentration was calculated again according to C1V1=
C2V2. For experiments studying cell membrane permeabilization
and trichome decay, the final biomass concentration was 1.5 g
l−1. For extraction of PC, the final biomass concentration was
set to 5 g l−1. The higher biomass for extraction was chosen to
have a higher concentration of PC in the sample. This allows for a
more accurate photometric measurement, as low concentrations
are close to the blank value. For better comparability, the PEF
treatment to measure particle decay was also performed at 5 gl−1
in this experiment.
Treatment Conditions
PEF treatments were performed in a prototype device designed
in-house. The exact configuration is described elsewhere
(Knappert et al., 2020). Voltage and electric current were directly
measured in the treatment chamber by a 75-MHz high-voltage
probe and a 100-MHz electric current probe, respectively. The
measured signals were visualized and stored with a digital
200 MHz oscilloscope (TDS2022B; Tektronix, United States).
Electroporation cuvettes (VWR) with a gap width of 4 mm and a
volume of 800 µl served as treatment chambers. For temperature
control during the experiment, the cuvettes and the PEF device
were placed below a temperature-controlled incubator hood
(Certomat, HK, Sartorius, Germany) for at least 30 min before
each experiment. The sample was preheated for 15 min to reach
the desired temperature using a thermomixer (Hettich Benelux
B.V., Netherlands) by shaking at 300 rpm.
In the first series of the experiments, the effect of electric field
strength, initial treatment temperature, and treatment time were
investigated. The applied treatment conditions are summarized
in Table 2. Exponential pulses were applied with a time constant
of 2 ±.63 µs and a pulse repetition rate f=1 Hz. The
pulse counts (number of applied pulses) for the various treatment
conditions were selected such that the critical parts of the
expected curve for cell permeabilization can be well-represented.
These are, namely, parts where cell disintegration rises and
reaches a plateau. After the treatment, each sample was diluted
to a suitable concentration for the following measurements (cell
membrane permeabilization for PI and particle size decay, see
following paragraphs) by transferring the treated sample to an
ice-cooled resuspension medium. The samples were stored on ice
until further analysis.
The specific energy input wspec of the treatment was calculated
as follows:
wspec =1
mnpZf−1
0
U(t)I(t)dt (1)
where mis the mass of the sample (800 µg, estimated density =
1 g cm−3), npis the number of applied pulses, and U(t) and I(t)
stand for voltage and electric current, respectively.
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Knappert et al. Phycocyanin Release PEF Arthrospira platensis
TABLE 2 | Number of pulses applied to the sample for different pulsed electric
field (PEF) treatments.
Initial treatment temperature [◦C]
Electric field
strength [kV cm−1]
30 40 50
5 6, 24, 40, 55,
65, 80, 90, 120
6, 24, 40, 55,
65, 80, 90, 120
6, 24, 40, 55,
65, 80, 90, 120
10 1, 6, 12, 24, 40,
55, 65, 70
1 ,6, 12, 24, 32,
40, 55, 65
1 ,6, 12, 24, 32,
40, 55, 65
15 1, 3, 6, 12, 24,
32, 40, 50, 60
1, 3, 6, 12, 24,
32, 40, 50, 60
1, 3, 6, 12, 24,
32, 45, 60
20 1, 2, 3, 6, 12,
24, 32, 40
1, 2, 3, 6, 9, 12,
18, 24, 32
1, 2, 3, 6, 9, 12,
18, 24, 32
25 1, 2, 3, 6, 12,
18, 24, 32
1, 2, 3, 6, 12,
18, 24, 28, 32
1, 2, 3, 6, 9, 12,
18, 24, 32
Analytical Methods
Measurement of Cell Permeabilization
To detect the effect of PEFs on permeabilization of the cell
membrane, a novel method based on fluorescence spectrometry
was developed in-house. PI, a substance that cannot penetrate
intact lipid membranes, was used as a dye. This substance
has already been used as a dye to detect cell disintegration of
phototrophic microorganisms (Luengo et al., 2014; Knappert
et al., 2020). For the development of the method, heat-
inactivated cells were used (80◦C, 15 min) to minimize pigment
interference with PI. The parameters biomass concentration,
PI concentration, staining time, and staining temperature
were varied based on experimental designs. After the optimal
staining parameters were found, the excitation and emission
wavelengths were adapted in order to measure samples with high
pigment content (i.e., PEF-treated samples). For a more detailed
description of the method development, refer to Appendix A.
Fluorescence measurements were conducted using a fluorescence
spectrofluorometer (RF-6000; Shimadzu, Japan).
The final method consists of the following steps. First, a PEF-
treated sample was diluted using an ice-cooled resuspension
medium to a biomass concentration of CB=0.25 g l−1,
and 1.8 ml of the diluted sample was mixed with 200 µl PI
stock solution. The final PI concentration in the sample was
cPI =11.2 µg ml−1. One unstained sample per treatment was
prepared to check whether cellular autofluorescence was affected
by the treatment, which was not the case for all the treatment
conditions (data not shown). The samples were shaken using
a thermomixer (Hettich Benelux B.V., Netherlands) at 23.1◦C
and 300 rpm for 17.5 min. After incubation, the sample was
transferred to a quartz cuvette (Helma Analytics, No. 101-10-
40, Germany) and measured directly at an emission wavelength
of 580 nm (bandwidth 5 nm) and an excitation wavelength of
490 nm (bandwidth 5 nm).
The degree of cell permeabilization was characterized by the
cell disintegration index ZDas defined in Lebovka et al. (2002):
ZD=Z−Z0
Zm−Z0(2)
Here, Zis the measured fluorescence signal, Z0is the fluorescence
signal of the untreated sample, and Zmis the maximum
fluorescence signal at complete cell disintegration. To obtain
Zm, reference treatment conditions were defined in pretrials at
which the fluorescence signal reached its maximum. Here, 25
kV cm−1and 32 pulses were chosen as reference treatment
conditions at a given temperature (cf. Table 2). These parameters
were validated by trying to recultivate the treated cells in a fresh
full SOT medium. After 3 days, the cells were agglomerated and
started to bleach out, and no growth occurred. Therefore, these
treatment conditions were considered to cause full irreversible
membrane permeabilization.
For all the treatment conditions (Table 2), the cell
disintegration index was determined in two biological replicates
and two technical replicates of the PEF treatment. Three
technical replicates were conducted for the reference treatment
and the untreated sample. The staining of each sample was
conducted in duplicate. Before calculating ZD, the arithmetic
mean of all four or six stained samples was calculated for each
biological repetition and inserted into Equation 2 for Z,Z0,
and Zm, respectively. For each treatment condition, we report
the arithmetic average of the two biological repetitions and
corresponding standard deviations.
Measurement of Cell Size Distribution
An analysis of cell size distribution after treatment was
performed using microscope images and a self-developed
image segmentation algorithm. Cells were treated with the
parameter combinations listed in Table 2, with the highest
applied pulse repetition number. Directly after the treatment,
the treated sample was diluted to a dry biomass concentration
of to CB=0.25 g l−1(OD750 =0.5). RGB images were
taken using a light microscope (Primo Star; Zeiss, Germany, 40×
magnification), equipped with a digital camera (Digital Sight DS-
U3; Nikon). The treated cell samples were tempered at 30◦C in
a thermomixer at 300 rpm. After 0, 30, 60, 120, and 180 min, a
sample was taken, and microscopic images of the cells were taken.
At least 700 cells were detected (untreated sample, 180 min).
Additionally, an untreated and a treated sample were kept at 30◦C
without shaking to investigate if the shaking itself has any impact
on cell size distribution. For image acquisition, a sample was
transferred to a counting chamber (Thoma counting chamber;
Hecht Assistant, Germany) and covered with an optical cover
glass. A total of 20 images were taken for each PEF treatment
and time at which samples were taken (0, 30, 60, 120, and 180
minutes).
The obtained images were processed using the commercial
software MATLABR2021a. The image processing code can
be obtained upon request to the corresponding author. The
algorithm is schematically shown in Figure 1. First, an RGB
image was converted into a grayscale. Then, the image was
smoothed using a Gaussian filter, and the contrast was enhanced
based on the standard deviation of the gray values. After
background subtraction, the image was converted into a binary
image. For this, we first selected upper and lower thresholds
based on grayscale histogram values to create two binaries, A
and B, of which we computed the intersection A∩(¬B). Objects
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