KR102222017B1 - Wild type and surface functional group modified microbial biofilm mediated nanoplastic removal method - Google Patents

Abstract

The present invention relates to a method for removing nanoplastics using a wild-type and surface functional group-modified microbial biofilm having nanoplastic aggregation and adsorption capacity,
The nanoplastic removal method according to the present invention has the effect of removing 99% or more of the nanoplastics present in the aqueous solution within a short time.

Description

{Wild type and surface functional group modified microbial biofilm mediated nanoplastic removal method}

The present invention relates to a method for removing nanoplastics using a wild-type and surface functional group-modified microbial biofilm having nanoplastic aggregation and adsorption capacity.

When plastic waste is exposed to nature, it is split into microplastics less than 5 mm by natural weathering. However, this cleaving process does not stop at the micro-scale, but proceeds further, breaking down to nanoplastics (< 1 μm). Microplastics have a number of problems, but how to get rid of them is not difficult. Filtering the water contaminated by microplastics can remove microplastics and purify them with clean water. However, nanoplastics are too small to be removed by filtering, and nanoplastics have different problems from microplastics because of their small size. Because nanoplastics are very small, they can cross biofilms, show toxicity, and are extremely difficult to detect and analyze. And because of this small size, it is also difficult to remove. Nanoplastics are plastics that are small enough to be difficult to observe even with an eye or a microscope, unlike microplastics that are several millimeters (< 5 mm). In addition, the effects of microplastics on the environment and organisms are known, but the specific absorption, distribution, and biological effects of nanoplastics in the body are not yet known. In this situation, there is a need for a method to remove nanoplastics.

Because nanoplastics have surface charges, there is an attractive force by van der Waals force and a repulsive force by electric charge between the nanoplastics. Because of this repulsive force, nanoplastics do not aggregate and exist in the form of dispersed nanoparticles. If the surface charge, which is the cause of this repulsive force, is canceled, the nanoplastics are aggregated or adsorbed. In other words, when the repulsive force is weakened due to the cancellation of the electric charge, it cannot exist as a nanoplastic in the first place because it aggregates. Nanoplastics have a wide variety of properties, and because various substances are adsorbed, their properties are very diverse. However, since this surface charge is a property that all nanoplastics have in common, a method to remove nanoplastics can be devised by using this property inversely.

Microbial biofilm is a entangled and shaggy aggregate of various types of organic matter (Non-Patent Document 1, PERIODICUM BIOLOGORUM Vol.109, No.2, 2007; Characteristics and significance of microbial biofilm formation), and various types of functional groups exist on the surface. . These functional groups have a different charge. The present inventors confirmed that the wild-type biofilm rapidly aggregated and adsorbed nanoplastics having a positive surface charge, thereby confirming the nanoplastic removal ability of the wild-type biofilm. However, the wild-type biofilm could not remove the surface negatively charged nanoplastics. In the biofilm, both negatively charged functional groups and positively charged functional groups exist, but it was hypothesized that positively charged functional groups interfere with the ability of positively charged functional groups to remove surface negatively charged nanoplastics because the negatively charged functional groups are more dominant. ^{Therefore, COO -} , a representative group of negatively charged functional groups, was subjected to esterification with strong acid and methanol to COOCH ₃ to prevent negative charges. This is called a surface functional group modified biofilm. This modified biofilm was confirmed that the surface negatively charged nanoplastics were rapidly agglomerated and adsorbed, resulting in removal capability. Thus, the surface positively charged nanoplastics were aggregated and adsorbed with the wild-type biofilm and the negatively charged nanoplastics were aggregated with the modified biofilm. Since the volume of the nanoplastic aggregates increased through this process, it was confirmed that more than 99% of the nanoplastics can be removed by applying filtering to remove them, thereby completing the present invention.

An object of the present invention is to provide a nanoplastic removal method capable of removing 99% or more of nanoplastics.

Another object of the present invention is to provide an adsorbent for removing nanoplastics.

Another object of the present invention is to provide a coagulant for removing nanoplastics.

Another object of the present invention is to provide a method of inducing agglomeration of nanoplastics to enable detection.

To achieve the above object,

One aspect of the present invention provides a nanoplastic removal method comprising the step of contacting a microbial biofilm with the nanoplastic.

In addition, another aspect of the present invention provides an adsorbent for removing nanoplastics including a microbial biofilm.

Further, another aspect of the present invention provides a coagulant for removing nanoplastics including a microbial biofilm.

Further, another aspect of the present invention provides a nanoplastic detection method comprising the step of contacting a microbial biofilm with the nanoplastic.

The nanoplastic removal method according to the present invention has the effect of removing 99% or more of the nanoplastics present in the aqueous solution within a short time.

1 is a diagram photographing a state in which a nanoplastic fluorescent Amine-PS having a positive surface charge is aggregated and adsorbed by a wild-type biofilm of a methylobacterium hispanicum strain.
Figure 2 is a graph comparing the size distribution of the Amine-PS aggregates aggregated by the wild-type biofilm compared to before putting the biofilm. In addition, it is a graph showing the change in the size distribution of the aggregates after vortexing and ultrasonication in order to show the resistance to the physical impact of the aggregates.
FIG. 3 is a diagram showing fluorescence photographing of fluorescent Amine-PS aggregates aggregated by a wild-type biofilm naturally sinking to the floor.
FIG. 4 is a diagram of fluorescence photographing that fluorescent Amine-PS is adsorbed onto a wild-type biofilm and thus the biofilm also exhibits fluorescence.
5 is a diagram photographed with an electron scanning microscope (SEM) of the aggregation and adsorption of Amine-PS on a wild-type biofilm.
6 is a diagram showing a method of modifying the surface functional groups of a wild-type biofilm.
7 is a diagram showing an infrared spectrum measured by FT-IR of a wild-type biofilm and a biofilm having a surface functional group changed.
Figure 8 is a graph measuring methanol produced by inducing ester-linked hydrolysis by putting a biofilm having a modified functional group in a strong base (0.25 M NaOH).
9 is a graph comparing surface charge changes by pH of a wild-type biofilm and a biofilm having a functional group modified.
10 is a diagram photographing a state in which a biofilm with a functional group modified and a fluorescent Sulfate-PS, a nanoplastic having a surface negative charge, are aggregated and adsorbed.
11 is a graph comparing the size distribution of Sulfate-PS aggregates agglomerated by the functional group-modified biofilm before the modified biofilm and after the wild-type biofilm. In addition, it is a diagram showing the change in size distribution after vortexing and ultrasonic treatment in order to show the resistance of the aggregates to mechanical impact.
FIG. 12 is a diagram of fluorescent Sulfate-PS aggregates naturally subsided.
13 is a diagram illustrating fluorescence imaging of a biofilm with a functional group modified to show fluorescence by adsorbing fluorescent Sulfate-PS.
14 is a diagram photographed by an electron scanning microscope (SEM) of a biofilm having a modified functional group on which Sulfate-PS is adsorbed.
FIG. 15 is a diagram showing a result of removing Amine-PS by filtering an aqueous Amine-PS solution aggregated and adsorbed with a wild-type biofilm through a membrane filter by fluorescence imaging.
16 is a graph showing the Amine-PS removal rate by reaction time by a method of aggregation and adsorption with a wild-type biofilm and then filtering.
FIG. 17 is a graph showing the Amine-PS removal rate by a method of aggregation and adsorption with a wild-type biofilm and then filtration by pH of an Amine-PS solution.
FIG. 18 is a diagram showing a result of removing Sulfate-PS by filtering an aqueous Sulfate-PS solution agglomerated and adsorbed with a functional group-modified biofilm through a membrane filter by fluorescence imaging.
19 is a graph showing the reaction of Sulfate-PS removal rate by a method of agglomeration and adsorption with a biofilm having a functional group modified and then filtering.
20 is a graph showing the Sulfate-PS removal rate by a method of agglomeration and adsorption with a modified biofilm and then filtration by pH of a Sulfate-PS solution.

Hereinafter, the present invention will be described in detail.

One aspect of the present invention provides a nanoplastic removal method comprising the step of contacting a microbial biofilm with the nanoplastic.

In this case, the microbial biofilm may be one or more biofilms selected from the group consisting of a wild type biofilm and a biofilm with modified surface functional groups. The wild type biofilm has a negative charge, so that nanoplastics having a positive charge can be aggregated and adsorbed, and the biofilm having a modified surface functional group has a positive charge, so that nanoplastics having a negative charge can be aggregated and adsorbed. have. That is, when the microbial biofilm is used in combination with a wild type biofilm and a biofilm modified with a surface functional group when removing nanoplastics, very excellent nanoplastic removal is possible regardless of the kind of charge that the nanoplastic stands out.

Meanwhile, the surface functional group-modified biofilm may be prepared by reacting a wild-type biofilm in the presence of an acid and an alcohol ^{to esterify the -COO-group present in the wild-type biofilm.} The biggest reason that the wild-type biofilm has a negative charge as a whole is ^{due to the -COO-} group present on the surface of the wild-type biofilm, and if esterification is performed, negative charges can be blocked. Accordingly, the biofilm whose surface functional groups have been modified by the positive charge existing in the wild-type biofilm can take on a positive charge as a whole.

The acid may be a known strong acid, preferably hydrochloric acid. The alcohol may be used without limitation as long as it is a known alcohol, but methanol may be preferably used.

The microbial biofilm may be a microbial biofilm of the genus Methylobacterium.

More specifically, the microorganisms of the genus Methylobacterium are M. hispanicum, M. adhaesivum, M. aerolatum, M. aminovorans, M. aquaticum, M. brachiatum, M. brachythecii, M. bullatum, M. cerastii , M. dankookense, M. extorquens, M. fujisawaense, M. gnaphalii, M. goesingense, M. gossipiicola, M. gregans, M. haplocladii, M. iners, M. isbiliense, M. jeotgali, M. komagatae, M. longum, M. marchantiae, M. mesophilicum, M. nodulans, M. organophilum, M. oryzae, M. oxalidis, M. persicinum, M. phyllosphaerae, M. phyllostachyos, M. platani, M. podarium, M. populi , M. pseudosasae, M. pseudosasicola, M. radiotolerans, M. rhodesianum, M. rhodinum, M. salsuginis, M. soli, M. suomiense, M. tardum, M. tarhaniae, M. thiocyanatum, M. thuringiense, M. trifolii, M. variabile, and M. zatmanii may be one or more selected from the group consisting of, preferably methylobacterium hispanicum (Methylobacterium hispanicum).

The nanoplastic may be a nanoplastic having an average diameter range of less than 1 μm. To take several average diameter ranges of nanoplastics to which the present invention can be applied, for example, it may be in the range of 1 to 900 nm, may be in the range of 1 to 800 nm, may be in the range of 1 to 700 nm, and in the range of 1 to 600 nm. May be, 1 to 500 nm range, 1 to 400 nm range, 1 to 300 nm range, 1 to 200 nm range, 1 to 100 nm range, 10 To 900 nm, 50 to 900 nm, 100 to 900 nm, 10 to 500 nm, 50 to 300 nm, and 70 to 200 nm May be, 80 to 150 nm range, 90 to 110 nm range, and 100 nm.

The nanoplastic may be a nanoplastic made of polystyrene, polyethylene, polypropylene, and polyethylene terephthalate.

The nanoplastic removal method may further include the step of inducing aggregation and adsorption of the nanoplastic by contacting the microbial biofilm with the nanoplastic, and then removing the nanoplastic through filtration.

In addition, the nanoplastic removal method may be to induce aggregation and adsorption of the nanoplastic by treating a microbial biofilm in an aqueous solution in which the nanoplastic is dispersed.

Another aspect of the present invention provides an adsorbent for removing nanoplastics including a microbial biofilm.

Another aspect of the present invention provides a coagulant for removing nanoplastics including a microbial biofilm.

At this time, since the detailed information on the microbial biofilm is the same as described above, it will be omitted to avoid redundant description.

Another aspect of the present invention provides a nanoplastic detection method comprising the step of contacting a microbial biofilm with the nanoplastic.

In the case of nanoplastics, their size is very small and it is generally difficult to detect. However, as in the present invention, when a microbial biofilm is brought into contact with the nanoplastics, aggregation of the nanoplastics is induced, so the detection of the nanoplastics is easy.

Hereinafter, the present invention will be described in detail through examples and experimental examples.

However, the examples and experimental examples to be described later are only to specifically illustrate the present invention in one aspect, and the present invention is not limited thereto.

Example 1. Confirmation of surface positively charged nanoplastic aggregation and adsorption using wild type biofilm of methylobacterium hispanicum strain

The stored methylobacterium hispanicum strain was streaked in TGY (Agar 1.5%) solid medium and grown at 30°C for 3 days until colonies emerged, and then only single colonies were picked. Put in a 14 ml tube containing 4 ml of TGY liquid medium (0.5% (w/v) tryptone, 0.1% (w/v) glucose and 0.3% (w/v) yeast extract) at 30° C. 200 rpm. It was pre-cultured for 3 days. Add 4 ml of TGY liquid medium to one well of a 6-well plate, add 400 µl of the pre-cultured biofilm solution, and incubate for 3 days at 30°C, pipette before inoculation to crush the biofilm of the pre-culture solution. I did. After 3 days, the formed biofilm was removed with tweezers and put into a new well containing 4 ml of distilled water, and the process of washing for 1 minute at 200 rpm was repeated three times. In a new well, add 3 ml of a 0.05% suspension obtained by diluting a 2.5% Amine-PS suspension 50 times in distilled water (Amine modified fluorescence polystyrene latex beads, 100 nm, sigma aldrich), and Amine-PS 0.05 contained in the well. % Suspension. After inserting the biofilm, it was shaken at 80 rpm to induce aggregation and adsorption, and then the change was observed.

1-1. Observation of surface positively charged nanoplastic aggregation and adsorption by biofilm

[Comparative Example 1]

Comparative Example 1-1. Suspension without added biofilm

Comparative Example 1-2. Add biofilm to distilled water

As a result, it was confirmed that agglomeration and adsorption occurred rapidly and particles were formed (see FIG. 1). Unlike Comparative Example 1-2, it was confirmed that adsorption occurred as the biofilm emit yellow light such as Amine-PS. Comparative Example 1-1 still emits yellow light, whereas in Example, yellow light is concentrated on the particles and biofilm by aggregation and adsorption, and it can be seen that fluorescence does not emit as much as distilled water in Comparative Example 1-2 in the solution. have. Therefore, it was confirmed that the nanoplastic contamination was purified by aggregation and adsorption of the nanoplastic by the biofilm.

1-2. Confirmation of the size of Amine-PS aggregates and their resistance to physical impact.

Dynamic light scattering (DLS) was used to confirm the size of the Amine-PS particles of the suspension in Comparative Example 1-1 and the size of the aggregates of Experimental Example 1. Since aggregation occurs in the external solution of the biofilm, the size of the aggregates can be determined by removing the biofilm with tweezers and collecting the supernatant containing aggregates and measuring by DLS. Compared to the Amine-PS of Comparative Example 1-1, the average volume of the Amine-PS aggregates of Example 1 was increased by ^{2×10 8 times.} In addition, the minimum diameter of the measured aggregate was 6560 nm, and there was no nanoplastic (< 1000 nm). In addition, vortexing 30 seconds and ultrasonication 30 seconds (130 watts, 20 kHz) were performed, respectively, to confirm the resistance to physical shock, and measured again by DLS. As a result, there was no significant change in the average size of the aggregate, and thus resistance to physical impact was confirmed (see FIG. 2).

1-3. Confirmation of Amine-PS sequestration and water purification by aggregation

It was confirmed that the Amine-PS of Example 1 aggregated and settled on the floor, and the supernatant was purified and did not exhibit fluorescence. On the other hand, it was confirmed that the Amine-PS suspension of Comparative Example 1 was not purified because fluorescence was evenly distributed throughout the solution. In Experimental Example 1-2, water purification was also confirmed in the case of vortexing and ultrasonic treatment (see FIG. 3).

1-4. Fluorescence confirmation of Amine-PS adsorption of biofilm.

In order to remove the fluorescence of the aggregates not adsorbed on the biofilm, the biofilm of Example 1 was removed with tweezers, placed in a new well containing distilled water, and washed at 200 rpm 3 times. The washed biofilm was put in fresh distilled water and subjected to fluorescence imaging, and the biofilm of Comparative Example 1-2 also went through the same process. Unlike Comparative Example 1-2, as fluorescence was observed throughout the biofilm, it was confirmed that not only aggregation but also adsorption occurred (see FIG. 4).

1-5. SEM confirmation of Amine-PS adsorption of biofilm.

The biofilms of Example 1 and Comparative Example 1-2 were observed with a scanning electron microscope (SEM). In Example 1, it was confirmed that nanoplastic particles, which were not observed in Comparative Example 1-2, were aggregated and adsorbed to the outer fibers of the cell. This suggests that the biofilm has the ability to adsorb surface positively charged nanoplastics (see Fig. 5).

Wild-type biofilm has the ability to remove surface positively charged nanoplastics (Amine-PS), but no such ability was found in the case of surface negatively charged nanoplastics (Sulfate-PS). (FIG. 10) Therefore, it was attempted to solve this by changing the surface functional group. Various kinds of functional groups exist in biofilm. Examples are COO ^- negatively charged and NH ₃ ^{+ positively charged.} The negatively charged functional group aggregates and adsorbs the surface positively charged nanoplastics through the attraction between the positive charge cancellation and the opposite charge. It was judged that the fact that the wild-type biofilm did not aggregate and adsorb the surface negatively charged nanoplastics is that the negatively charged functional groups exist more predominantly than the positively charged functional groups on the surface of the biofilm, thereby inhibiting the aggregation and adsorption of the positively charged functional groups of the surface negatively charged nanoplastics. Therefore, in order to block the charge of the negatively charged functional group, a change in the surface functional group was induced. When the biofilm is put in methanol containing 0.1 M HCl and reacted at 65° C. for 12 hours, the surface functional group R-COO ^- is changed to R-COOCH _3. R-COO ^- has a negative charge, whereas R-COOCH ₃ has no charge. Therefore, the strong negative charge on the surface of the biofilm is canceled, and the surface negatively charged nanoplastics can be aggregated and adsorbed. (Fig. 6)

Example 2-1. Confirmation of surface functional group change through FT-IR specta comparison.

25 ml of TGY liquid medium was added to the Petri dish, and 500 μl of the pre-culture solution was added. A total of 10 were prepared. After incubation at 30° C. for 3 days, all 10 biofilms formed were removed with tweezers and washed with DW. A 50 ml Conical tube was filled with 40 ml of methanol containing 0.1 M HCl. All washed biofilms were removed with tweezers and placed in a conical tube. The reaction was carried out at 65° C. for 12 hours to induce surface functional group modification. After the reaction, the Conical tube was centrifuged at 4000 rpm for 1 minute, the supernatant was discarded, and methanol was even filled up to 50 ml for washing and rotated at 200 rpm for 5 minutes. Centrifuged again at 4000 rpm for 1 minute, discarded the supernatant, this time filled to 50 ml with DW, and shaken at 200 rpm for 5 minutes. After washing twice with DW and discarding the supernatant, quick freezing at -80°C for 1 hour and then freeze-dried overnight at -65°C. And FT-IR was measured.

Comparative Example 2-1. Wild-type biofilm without surface functional group change reaction

As a result, three peaks corresponding to the constituents of a new ester bond that were not present in Comparative Example 2-1 were confirmed. In addition, the peak of 1440 cm-1 was smaller than the right neighboring peak in Comparative Example 2-1, but it was confirmed that it was larger than the right neighboring peak in Example 2-1. This is a peak corresponding to (C-H), meaning that the ratio of C-H has increased. This indicates that the intended surface functional group, the methyl ester bond, was increased.

Example 2-2. Confirmation of the amount of change in surface functional groups

When the surface functional group-modified biofilm is added to strong base water, a hydrolysis reaction is induced to separate R-COOCH3 into R-COOH and methanol again. Therefore, when the biofilm having the surface functional group modified is added to a strong base and reacted and the resulting methanol is measured, it can be confirmed that the ester bond is formed by methanol, that is, the surface functional group modification has occurred, and the degree can be quantified. To confirm this, 50 ml of TGY liquid medium and 500 μl of the pre-cultured solution were added to two 250 ml Erlenmeyer flasks, respectively, and incubated at 30° C. for 4 days. This culture solution was placed in a 50 ml conical tube and centrifuged at 4000 rpm for 1 minute. The supernatant was discarded, another culture solution was added, centrifuged at 4000 rpm for 1 minute, and the supernatant was discarded. Filled up to 40 ml with distilled water, and shaken for 5 minutes at 200 rpm for washing. Centrifugation was performed at 4000 rpm for 1 minute, and the supernatant was discarded and washed with distilled water. After discarding the supernatant, -80 ℃ rapid freezing was carried out for 1 hour and freeze-dried overnight at -65 ℃.

5 ml of methanol containing 0.1 M HCl was added to a 50 ml conical tube containing lyophilized wild-type biofilm, and shaken vigorously, rotated at 200 rpm for 5 minutes, and reacted at 65° C. for 12 hours. After that, the washing process was repeated 5 times in which the conical tube was filled with DW up to 50 ml, rotated at 200 rpm for 5 minutes, and centrifuged at 4000 rpm for 5 minutes. The last centrifugation was performed at 4000 rpm for 10 minutes. The supernatant was discarded and freeze-dried overnight at -65°C after 1 hour of rapid freezing at -80°C. After weighing the freeze-dried surface functional group-modified biofilm, it was transferred to a new 15 ml conical tube, and 0.25 M NaOH of weight×(ml/mg) was added. This was vortexed and mixed strongly, and then shaken at 200 rpm for 30 minutes, and then overnight at 4°C. Then, after centrifugation at 4000 rpm for 10 minutes, the supernatant was filtered with a 0.2 μm CA syringe filter. 1 mM of 1-butanol was added to the filtered solution as an internal standard, and the amount of methanol was measured using GC/FID (see FIG. 8). (GC/FID condition: Injector temp 250℃, column temp 40℃, detector temp 270℃, run time 5 min, 1:100 split)

[Comparative Example 2]

Comparative Example 2-2-1. Distilled water was added instead of 0.25 M NaOH in Example 2-2

Comparative Example 2-2-2. In Example 2-2, before modification of the surface functional group, that is, 0.25 M NaOH was added to the wild-type biofilm and then the same process was performed.

Methanol was not detected in any of the comparative examples, and methanol was detected only in Example 2-2. As a result of Comparative Example 2-2-1, doubts about the reliability of the experiment, such as that the washing before the hydrolysis reaction was not properly carried out, and residual methanol remained, can be eliminated, and that the hydrolysis by the strong base was performed correctly. Able to know. In Comparative Example 2-2-2, since methanol was not detected, it could be confirmed that there was no or very little methyl ester on the surface of the wild-type biofilm. Therefore, it can be confirmed that methanol entered the surface functional group of the biofilm in the form of methyl ester by the modification of the surface functional group.

Example 2-3. Confirmation of biofilm surface charge change due to surface functional group change.

After incubating for 3 days at 30° C. with 4 ml of TGY and 400 μl of pre-culture in a 6 well palte, the cultured wild-type biofilm was washed with distilled water. In a 1.5 ml microtube, 1 ml of methanol containing 0.1 M HCl and the washed biofilm were placed, and reacted at 65° C. for 12 hours to induce a change in surface functional groups. The biofilm having the surface functional group changed was taken out with tweezers and washed sequentially with methanol and distilled water. 100 μl of distilled water and the washed modified biofilm were put in a 96 well plate, and then rapidly frozen at -80°C and freeze-dried at -65°C. Freeze-dried modified biofilm was added to 1 ml of distilled water of pH 2, 4, 6, 8, 10, 12, respectively, pulverized by pipetting, and the Zeta potential was measured with ELSZ-1000 (Otsuka, japan).

Comparative Example 2-3. Zeta potential measurement of wild-type biofilm.

As a result, the wild-type biofilm of Comparative Example 2-3 exhibits a stable negative charge in the pH range of 4 to 12, while the surface functional group-modified biofilm of Example 2-2 exhibits a stable positive charge in the pH range of 2 to 10. It was confirmed that electrostatic properties were changed due to functional group charge blocking (see FIG. 9).

Confirmation of agglomeration and adsorption of negatively charged surface nanoplastics using biofilm modified with surface functional groups

4 ml of TGY and 400 μl of pre-culture were added to a 6 well plate, and the wild-type biofilm grown at 30° C. for 3 days was washed with distilled water. 1 ml of methanol containing 0.1 M HCl was added to a 1.5 ml microtube, and the washed biofilm was put in with tweezers. This was reacted at 65° C. for 12 hours, and then washed with methanol and water. In a new 6 well, 3 ml of a 0.05% suspension diluted 50 times in distilled water was added to a 2.5% sulfate-PS suspension of surface negatively charged nanoplastic (Sulfate modified fluorescence polystyrene latex beads, 100 nm, sigma aldrich). Three biofilms with functional group modification were added to the 0.05% suspension contained in the well and shaken at 80 rpm to induce aggregation and adsorption. It was confirmed that agglomeration and adsorption occurred from 1 minute after starting the reaction to form particles (see FIG. 10).

[Comparative Example 3]

Comparative Example 3-1 Only the same suspension as in Example 3 was rotated at 80 rpm.

Comparative Example 3-2 The wild-type biofilm of Example 1 was added to the same suspension as in Example 3 and rotated at 80 rpm.

Comparative Example 3-3 Biofilm with changed surface functional groups added to distilled water

3-1. Observation of surface negatively charged nanoplastic aggregation and adsorption by surface functional group-modified biofilm

Unlike Comparative Example 3-3, it was confirmed that adsorption occurred as the biofilm emit pink light such as Sulfate-PS. Comparative Example 3-1 and Comparative Example 3-2 still emit pink light, whereas in Example 3, pink light was concentrated on the particles and the biofilm by aggregation and adsorption, and the fluorescence of Comparative Example 3-3 was concentrated in the solution. It was confirmed that it did not emit light as much as distilled water. Therefore, it was confirmed that it was purified from nanoplastic contamination by aggregation and adsorption of the nanoplastic by the biofilm.

3-2. Confirmation of the size of sulfate-PS aggregates and their resistance to physical impact.

Dynamic light scattering (DLS) confirmed the size of the Sulfate-PS particles of the suspension of Comparative Example 3-1 and the size of the aggregates of Example 3. Since aggregation occurs in the external solution of the deformed biofilm, the size of the aggregates can be determined by removing the deformed biofilm with tweezers and collecting the supernatant containing aggregates and measuring DLS. Compared to the Sulfate-PS of Comparative Example 3-1, the average volume of the Sulfate-PS aggregates of Example 3 was ^{increased by 5×10 9} times or more. In addition, the minimum diameter of the measured aggregate was 2379.1 nm, and there was no nanoplastic (< 1000 nm). In order to check the resistance to physical shock, vortexing was performed for 30 seconds and ultrasonication was performed for 30 seconds (130 watts, 20 kHz), respectively, and DLS was measured again. As a result, there was no significant change in the average size of the aggregate, and thus resistance to physical impact was confirmed (see FIG. 11).

3-3. Sulfate-PS sequestration by flocculation and confirmation of water purification

It was confirmed that the Sulfate-PS of Example 3 aggregated and settled on the floor, and the supernatant was purified and did not exhibit fluorescence. On the other hand, it was confirmed that the fluorescence of the Sulfate-PS suspensions of Comparative Examples 3-1 and 3-2 were evenly distributed throughout the solution and were not purified at all. Water purification was also confirmed in the case of vortexing and ultrasonic treatment in Experimental Example 3-2 (see FIG. 12).

3-4. Fluorescence confirmation of Sulfate-PS adsorption of the surface functional group-modified biofilm.

In order to remove the fluorescence of the aggregates not adsorbed on the biofilm, the biofilm of Example 3 was removed with tweezers, placed in a new well containing distilled water, and washed at 200 rpm 3 times. The washed biofilm was put again in fresh distilled water and subjected to fluorescence imaging, and the biofilm of Comparative Example 3-3 also went through the same process. Unlike Comparative Example 3-3, as fluorescence was observed throughout the biofilm, it was confirmed that not only aggregation but also adsorption occurred (see FIG. 13).

3-5. SEM confirmation of Sulfate-PS adsorption of the modified biofilm.

The biofilms of Example 3 and Comparative Examples 3-2 and 3-3 were observed with a scanning electron microscope (SEM). In Example 2, it was confirmed that nanoplastic particles, which were not observed in Comparative Examples 3-2 and 3-3, were aggregated and adsorbed on external fibers. Therefore, it was confirmed that the modified biofilm has the ability to adsorb surface negatively charged nanoplastics (see FIG. 14).

Removal of nanoplastics by filtering after aggregation and adsorption using biofilm

4-1. Amine-PS coagulation and adsorption with wild-type biofilm and filtering to remove

Since the size of the nanoplastic increased due to the aggregation and adsorption of the nanoplastic using the biofilm, it was removed by filtering. 1 ml of Amine-PS 0.05% suspension and biofilm were added to a 6 well plate, and reacted at 120 rpm for 15 minutes. A 1 ml syringe was mounted on a PVDF syringe filter (pore size 0.2 μm, hyundai micro), and the agglutination solution was filtered. The edge of the filter was cut with scissors, the filter membrane inside was taken out, and dried for one day. This was emitted with Epi-blue (460-490 nm) with a Bio-rad Chemi-Doc MP imaging system, and fluorescence was photographed using a 590/110 nm standard filter. Compared to the control group in which the biofilm was not added, the filter of the experimental group showed strong fluorescence, and the filtrate did not show fluorescence, so that Amine-PS was caught in the filter and the water was purified. On the other hand, it was confirmed that the filter of the control group had very weak fluorescence, and the filtrate showed strong fluorescence, so that almost no water purification occurred.

4-2. Amine-PS removal rate by reaction time with wild-type biofilm

In order to check the removal rate of wild-type biofilm over time, 4 ml of TGY and 400 ul of pre-culture solution were added to a 6 well plate, and two biofilms cultured at 30° C. for 3 days were washed with distilled water. 1.5 ml of Amine-PS 0.05% suspension and the washed biofilm were added to 6 wells, and reacted at 120 rpm. After the reaction was started, 150 μl each was sampled 30 seconds, 1 minute, 5 minutes, 15 minutes, and 1 hour, and filtered with a PVDF syringe filter. The control group filtered the Amine-PS suspension rotated at 120 rpm for 1 hour without biofilm. n=3. As a result, it was confirmed that the removal rate exceeded 99% in 1 minute of reaction time (see FIG. 16).

4-3. Amine-PS removal rate according to pH change

In a 12 well plate, add 2 ml of TGY and 200 ul of the pre-culture solution, wash the biofilm cultured at 30°C for 3 days with distilled water, and then add it to 96 wells containing 100 μl of distilled water in advance, and rapidly at -80°C. Freeze-dried after freezing. Distilled water having a pH of 2, 4, 6, 8, and 10 was prepared by adjusting the pH with 1 M of NaOH and 1 M of HCl in distilled water. A 0.05% Amine-PS suspension for each pH was prepared by diluting 2.5% Amine-PS 50 times in distilled water adjusted to pH. 150 μl of a pH-adjusted 0.05% Amine-PS suspension was added to 96 wells containing the lyophilized biofilm and rotated at 300 rpm for 15 minutes. After 15 minutes, 150 μl was filtered with a PVDF syringe filter. The control was rotated for 15 minutes at 300 rpm without biofilm and filtered. As a result, it was confirmed that the removal rate was 99% or more in the range of pH 4-10 (n=3).

4-4. Sulfate-PS coagulation and adsorption with a biofilm with a changed surface functional group, followed by filtering to remove

1 ml of Sulfate-PS 0.05% suspension was added to a 6 well plate, and the biofilm grown in a 6 well plate and modified surface functional group was added and reacted at 120 rpm for 15 minutes. A 1 ml syringe was mounted on a CA syringe filter (pore size 0.2 μm, hyundai micro) and the coagulation solution was filtered. The edge of the filter was cut with scissors, the inner filter membrane was taken out and dried for one day. This was emitted with Epi-blue (460-490 nm) with a Bio-rad Chemi-Doc MP imaging system, and fluorescence was photographed using a 590/110 nm standard filter. Compared to the control group in which no biofilm or wild-type biofilm was added, the filter of the experimental group emit strong fluorescence, and the filtrate did not emit fluorescence, so it was confirmed that all of the Sulfate-PS was caught in the filter and the water was purified. On the other hand, it was confirmed that the filter of the control group had very weak fluorescence, and the filtrate emitted strong fluorescence, so that almost no water purification occurred. (See Fig. 18)

4-5. Sulfate-PS removal rate by reaction time of biofilm with changed surface functional group

In order to check the time-dependent removal rate of the biofilm having a changed surface functional group, 4 ml of TGY and 400 ul of the pre-culture solution were added to a 6 well plate, and the biofilm cultured at 30° C. for 3 days was washed with distilled water. 1 ml of methanol containing 0.1 M HCl was added to a 1.5 ml microtube, and the washed biofilm was taken out with tweezers, reacted at 65° C. for 12 hours, and washed with methanol and distilled water. 100 μl of distilled water and a biofilm with changed surface functional groups were put in a 96 well plate, and then rapidly frozen at -80°C, and freeze-dried at -65°C. Two lyophilized biofilms with changed surface functional groups were put into a 6 well plate, 1.5 ml of a 0.05% Sulfate-PS suspension was added in the same well, and reacted at 200 rpm. 150 μl each was sampled 30 seconds, 1 minute, 5 minutes, 15 minutes, and 1 hour after the start of the reaction, and filtered with a CA syringe filter. The control group was a suspension rotated at 120 rpm for 1 hour without biofilm (n=3). As a result, it was confirmed that the removal rate exceeded 99% during the reaction time of 30 seconds (see FIG. 19).

4-6. Sulfate-PS removal rate according to pH change

In a 12 well plate, add 2 ml of TGY and 200 ul of the pre-culture solution, wash the biofilm cultured at 30°C for 3 days with distilled water, and put it in a 1.5 ml microtube containing 1 ml of methanol containing 0.1 M HCl, and at 65°C. It was reacted for 12 hours. After the reaction, it was washed with methanol and distilled water, added to 96 wells in which 100 μl of distilled water was put in advance, and freeze-dried after rapid freezing at -80°C. Distilled water of pH 4, 6, 8, and 10 was prepared by adjusting the pH with 1 M of NaOH and 1 M of HCl in distilled water. A 0.05% Sulfate-PS suspension for each pH was prepared by diluting 2.5% Sulfate-PS 50 times in distilled water whose pH was adjusted. 150 μl of a pH-adjusted 0.05% Sulfate-PS suspension was added to 96 wells containing a biofilm with a lyophilized surface functional group changed, and rotated at 300 rpm for 15 minutes. After 15 minutes, 150 μl was filtered with a CA syringe filter, and the fluorescence of the filtrate was measured to calculate the removal rate. The control group rotated and filtered for 15 minutes at 300 rpm without biofilm, and reacted with wild-type biofilm instead of biofilm with a surface functional group changed. As a result, it was confirmed that the removal rate was 99% or more in the range of pH 4-10 (n=3).

Claims (15)

It is a nanoplastic removal method comprising the step of contacting a microbial biofilm with the nanoplastic,
The microbial biofilm is a biofilm having a modified surface functional group,
The surface functional group-modified biofilm has a positive charge, characterized in that the agglomeration and adsorption of the nanoplastic having a negative charge, nanoplastic removal method.
The method of claim 1,
The microbial biofilm further comprises a wild type biofilm.
The method of claim 2,
The wild type biofilm has a negative charge, characterized in that the agglomeration and adsorption of the nanoplastic having a positive charge, nanoplastic removal method.
delete The method of claim 1,
The surface functional group-modified biofilm,
A method for removing nanoplastics, characterized in that produced by reacting a wild-type biofilm in the presence of an acid and an alcohol, ^{and esterifying the -COO-group present in the wild-type biofilm.}
The method of claim 5,
The acid is hydrochloric acid;
The alcohol is methanol, characterized in that, nanoplastic removal method.
The method of claim 1,
The microbial biofilm is a method of removing nanoplastics, characterized in that the microbial biofilm of the genus Methylobacterium.
The method of claim 1,
The nanoplastic is characterized in that the nanoplastic having an average diameter range of less than 1μm, nanoplastic removal method.
The method of claim 1,
The nanoplastic is characterized in that at least one selected from the group consisting of polystyrene, polyethylene, polypropylene, and polyethylene terephthalate.
The method of claim 1,
The nanoplastic removal method,
After inducing the aggregation and adsorption of the nanoplastic by contacting the microbial biofilm with the nanoplastic, the method further comprising removing the nanoplastic through filtration.
The method of claim 1,
The nanoplastic removal method,
A method for removing nanoplastics, characterized in that by treating a microbial biofilm in an aqueous solution in which the nanoplastics are dispersed to induce aggregation and adsorption of the nanoplastics.
The method of claim 11,
The nanoplastic removal method,
A method for removing nanoplastics, further comprising the step of inducing agglomeration of the nanoplastics and then precipitating them.
As a microbial biofilm, an adsorbent for removing nanoplastics comprising a microbial biofilm, characterized in that the surface functional groups are modified to have a positive charge and agglomerate and adsorb nanoplastics having a negative charge.
As a microbial biofilm, a coagulant for removing nanoplastics comprising a microbial biofilm, characterized in that the surface functional groups are modified to have a positive charge and agglomerate and adsorb nanoplastics having a negative charge.
As a microbial biofilm, comprising the step of contacting a microbial biofilm with a nanoplastic, characterized in that the surface functional group is modified to have a positive charge and a nanoplastic having a negative charge and agglomeration and adsorption, nanoplastic detection Way.

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