Lensless digital holographic microscopy as an efficient method to monitor enzymatic plastic degradation

https://doi.org/10.1016/j.marpolbul.2020.111950Get rights and content

Highlights

  • Enzymatic degradation of plastic film was analyzed via digital holographic microscopy.

  • Lensless microscopy is capable to monitor PET degradation.

  • Non-destructive testing of biological plastic degradation is possible via this method.

  • PET degradation by recombinant PETase was demonstrated over a time course of 43 days.

Abstract

A big challenge of the 21st century is to cope with the huge amounts of plastic waste on Earth. Especially the oceans are heavily polluted with plastics. To counteract this issue, biological (enzymatic) plastic decomposition is increasingly gaining attention. Recently it was shown that polyethylene terephthalate (PET) can be degraded in a saltwater-based environment using bacterial PETase produced by a marine diatom. At moderate temperatures, plastic biodegradation is slow and requires sensitive methods for detection, at least at initial stages. However, conventional methods for verifying the plastic degradation are either complex, expensive, time-consuming or they interfere with the degradation process. Here, we adapt lensless digital holographic microscopy (LDHM) as a new application for efficiently monitoring enzymatic degradation of a PET glycol copolymer (PETG). LDHM is a cost-effective, compact and sensitive optical method. We demonstrate enzymatic PETG degradation over a time course of 43 days employing numerical analysis of LDHM images.

Introduction

Plastics are synthetic polymers that have been intensively produced and advanced since the middle of the 20th century. In 2018, more than 350 million tons of plastics were synthetized – the majority from fossil feedstocks – and production rates are expected to further increase dramatically (plasticseurope.org). A significant portion of newly produced plastics is composed of thermoplastic polymers including polyethylene terephthalate (PET), polypropylene (PP), polyethylene (PE), polyvinyl chloride (PVC), and polystyrene (PS) (Chamas et al., 2020). The many beneficial properties of plastics, including durability, stability and flexibility, make these materials highly valuable for a wide variety of applications. For example, they are intensely used as packaging materials for food and beverages, synthetic clothing fibers, construction materials, and are also essential for many medical and technical products and processes. The development of plastic materials with a multitude of unique and industrially desirable characteristics has, without a doubt, facilitated and accelerated our technological progress in many ways.

Besides the many beneficial aspects of synthetic polymers, the current use of the majority of plastic products is far from being sustainable. As a consequence, gigantic amounts of plastic waste are produced incessantly, posing a huge challenge for waste management systems worldwide. Although effective industrial recycling processes for many plastic materials are established, a huge fraction of plastic waste is still incinerated or landfilled and due to improper waste disposal, mismanagement or heedless release, it ends up in the environment. Plastic pollution has become a serious issue in the last decades with so far unforeseeable consequences for Earth's ecosystems. Especially the oceans are polluted massively by an increasing entry of synthetic polymers, at a rate estimated about 4.8 to 12.7 million tons each year (Jambeck et al., 2015). Also, the strong increase of microparticles in marine environments has become a global issue that requires attention (Andrady, 2011).

As an ecofriendly alternative for existing industrial recycling processes for plastic waste as well as for bioremediation of environments polluted with plastic particles, an efficient biological degradation of plastics via microorganisms/enzymes is highly desirable. However, there are only very few microbial organisms known to be able to degrade petrochemical-based plastics. Especially PP, PE, PET, PS and PVC usually have an extremely limited biodegradability (see, e.g., (Shah et al., 2008; Urbanek et al., 2018; Danso et al., 2019)). This is mostly due to their synthetic/non-natural character and the exceptional stability of their chemical bonds as well as e.g. crystalline polymer structures (Wei and Zimmermann, 2017). These features make many plastics almost impervious to enzymatic degradation. Nevertheless, several microbial enzymes capable of plastic degradation have been discovered in recent years (see, e.g., (Danso et al., 2019) for review). One of these enzymes is PETase, a bacterial PET hydrolase, that disassembles PET into monomers, including mono(2-hydroxyethyl) terephthalic acid (MHET) and terephthalic acid (TPA) at moderate temperatures around 30 °C (Yoshida et al., 2016). The worldwide annual production rate of PET (for bottles, synthetic fibers, packaging, etc.) is predicted to exceed 70 million tons in 2020 (see (Palm et al., 2019) and references therein) making the polymer one of the most frequently used polyester-based thermoplasts on Earth.

Another enzyme, leaf-branch compost cutinase (LCC), can also degrade PET albeit at higher temperatures between 50 and 80 °C (Sulaiman et al., 2012). Recently, an LCC degrading PET into TPA and ethylene glycol (EG) has been further improved by bioengineering, and might have potential for large-scale industrial PET recycling processes (Tournier et al., 2020). However, LCC requires high temperatures (around 70 °C) for polymer hydrolysis, whereas PETase can degrade PET even at moderate temperatures, although at lower rates (Yoshida et al., 2016; Wei et al., 2019b). Thus, PETase is currently the enzyme of choice for PET bioremediation purposes in the moderate temperature range, whereas LCC is better suited for industrial PET degradation requiring conditions near the PET glass transition temperature (~75 °C) for efficient depolymerization.

Employing synthetic biology, we recently developed a transgenic marine diatom (Phaeodactylum tricornutum) to produce an engineered PETase (AP_SP-PETaseR280A-FLAG, see below) that is excreted by the cells and capable of PET degradation in saltwater environments at moderate temperatures (Moog et al., 2019). This algal cell factory holds potential for applications in bioremediation of seawater polluted with PET micro- and nanoparticles in a closed system such as a large-scale bioreactor or marine wastewater plant. Besides being active on industrially shredded PET and PET film from plastic bottles, the PETase produced by the microalga can also degrade the PET co-polymer PET glycol (PETG), a highly amorphous polymer consisting of dimethyl terephthalate, ethylene glycol und 1,4 cyclohexane dimethanol (Moog et al., 2019).

Biological degradation of plastic films is usually measured via one of the following methods: weight loss determination, scanning electron microscopy (SEM), atomic force microscopy (AFM), nuclear magnetic resonance (NMR) spectroscopy, Fourier-transform infrared (FTIR) spectroscopy, differential scanning colorimetry (DSC), high-performance liquid chromatography (HPLC) or a combination of two or more of these techniques (see, e.g., (Ioakeimidis et al., 2016; Yoshida et al., 2016; de Castro et al., 2017; Han et al., 2017; Ojha et al., 2017; Schmidt et al., 2017; Austin et al., 2018; Joo et al., 2018; Delacuvellerie et al., 2019; Moog et al., 2019; Wei et al., 2019a; Wei et al., 2019b; Tournier et al., 2020) and see (Shah et al., 2008) for additional methods). Raman spectroscopy is another tool for identification of plastics and measuring structural alterations of plastic film surfaces which may be caused by weathering or even biological degradation (Araujo et al., 2018; Dong et al., 2020). Whereas some of these methods including, e.g. HPLC, allow for quantification of the degradation products released by enzymatic reactions, structural alterations – for example surface degradation – of the substrates can be observed via well-established techniques such as SEM, AFM or, if distinct enough, even by confocal or conventional light microscopy. However, SEM is by far the most common microscopy method to analyze surface alterations of enzymatically treated plastic films. Although a huge benefit of SEM is the very high resolution for sample analysis up to the low nanometer range, one major disadvantage of SEM is that the sample surface for visualization has to be coated with a conductive material such as gold, thereby preserving the sample but preventing a further use of the plastic sample for additional (degradation) experiments. For these reasons, it would be highly desirable that the plastic material remains accessible and intact after visual analysis, allowing further processing and to follow the progress of degradation of a single sample, e.g., in a long-term experiment. Furthermore, the possibility of directly monitoring the plastic degradation for example for different reaction conditions and having immediate access to the results would be very useful. For this purpose, in addition to the conventional techniques, alternative methods have to be explored.

Measuring a sample without destroying or altering it is referred to as non-destructive testing (Shull, 2002). Optical metrology methods are contactless and thus well-suitable for non-destructive testing. The most common optical method is probably conventional light microscopy. However, these microscopes require sophisticated lens systems, which can be very expensive, while their usefulness is limited. Although they can measure effects like absorption and reflection very well, the contrast for transparent samples is often too low. Since plastic is mainly transparent or semitransparent, a measurement technique able to image phase objects and being sensitive to optical path length changes might be more suitable to monitor plastic surface degradation.

A well-established method with various applications in biology or materials research is Digital Holography (Kemper and von Bally, 2008; Kim, 2010, Edwards et al., 2014). It can produce fast single-shot images, provides amplitude as well as phase information and allows 3D topographic imaging. In Digital Holography an object light wave, which is altered by the sample, superimposes with a reference plane wave, which is unaltered. The resulting interference pattern is recorded by a digital camera and numerically reconstructed with the angular spectrum method to create amplitude, phase and 3D information. Furthermore, the reconstruction of holograms includes the possibility of numerical refocusing, so that samples do not need to be in focus to produce sharp reconstructed images in the end (Kim, 2011).

Commercial systems for holographic measurements are available and can for example be used for investigating electrochemical deposition processes (Abbott et al., 2013) and cell measurements (Hellesvik et al., 2020; Kamlund et al., 2017). For investigating plastic degradation, a stable holographic system is necessary. Among others the system HoloTop (Fratz et al., 2019) proved to provide reliable imaging even in very unstable industrial environments. However, these systems are expensive and not compact. In-line holography typically provides a stable setup and one very robust, cost-effective and simple setup is lensless digital holographic microscopy (LDHM) (Greenbaum et al., 2012). It only needs a laser diode for illumination, a place to fix the sample and a sensor to capture the diffraction pattern, which can be numerically reconstructed afterwards (Adinda-Ougba et al., 2015). LDHM is often used for biological applications or in low-resource settings (Xu et al., 2001; Moscelli et al., 2011; Ozcan and McLeod, 2016). Since it provides an uncomplicated imaging method, it can also be used by non-technical experts. Due to its few components, it is very insensitive to changes in the surrounding and thus always provides the same measurement conditions. For measurements in the marine environment, there are already approaches with digital holography to measure and identify microparticles (Merola et al., 2018; Grant-Jacob et al., 2019; Bianco et al., 2020) and plankton (Watson et al., 2001; Dyomin et al., 2020). Thus, this metrology technique can be easily adapted to the application and in contrast to the commercially available systems also holds potential for monitoring plastic degradation in saltwater.

Instead of performing numerical reconstruction of the diffraction pattern, it is also possible to analyze only the shadow image. For shadow imaging, the diffraction pattern captured by the sensor is not further reconstructed. It shows the shadows of the sample structures. Due to the missing numerical refocusing the structures are not sharp. Nevertheless, the similarity to the original structures is sufficient to detect and analyze changes of the sample (Ozcan and McLeod, 2016). Thus, shadow imaging provides the advantage of saving computational power and thus simplifies and accelerates the process further. Shadow imaging was already used, e.g., for monitoring cells (Jin et al., 2012). Overall, LDHM is a fast and cost-effective method that, due to its simple setup (few, cheap components) and adaptability, might be generally useful for in situ measurement of biologically caused plastic polymer decay – that is, monitoring a successive enzymatic degradation of a plastic film in real time. Thus, this technique might have individual advantages compared to other currently used visual methods for monitoring (biological) plastic degradation, such as SEM, for which the samples have to be “destroyed” (see above). However, whether a comparable degree and depth of information of (real-time) LDHM analysis compared to SEM could be achieved is unknown.

In this paper, we apply LDHM as a simple, stable and fast method for monitoring enzymatic plastic degradation using PETase as biocatalyst and PETG film as substrate. Furthermore, we show proof of the degradation by numerical analysis of the images. Afterwards we compare the results to the outcomes of the well-established methods SEM and ultra HPLC (UHPLC).

Section snippets

Enzyme synthesis

For time course enzymatic plastic degradation assays the bacterial PET hydrolase PETase (Yoshida et al., 2016) was produced using the eukaryotic microalga Phaeodactylum tricornutum as described earlier (Moog et al., 2019). In brief, a microalgal strain expressing a recombinant, activity enhanced PETaseR280A version (Joo et al., 2018) equipped with the signal peptide of a P. tricornutum alkaline phosphatase (AP_SP: first 30 amino acids of Phatr2 49678) at the N-terminus for efficient secretion

Results

Data was generated using the experimental setup as follows: First, PETase enzyme was synthesized by using the microalga P. tricornutum as a microbial cell factory and secreted into the saltwater medium of the culture (pH 8.0, 1.66% salinity). The PETase-containing culture medium was separated from the algal cells and used for plastic degradation experiments as previously described in (Moog et al., 2019). PETG film particles were incubated with the culture medium fractions of a PETase expressing

Discussion

Monitoring biological plastic degradation over a time course can be technically challenging. In case of PET, especially at moderate temperatures the process of enzymatic plastic breakdown is rather slow. Consequently, the change of the surface structure of the plastic is also rather gradual, making a sensitive detection method for surface alterations during enzymatic plastic degradation highly valuable. In addition, some of the commonly used analysis methods (such as SEM) require fixation of

Conclusion

Overall, LDHM is a useful and practical tool to verify the enzymatic plastic degradation of PETG film, especially in the initial stages of biodegradation. LDHM can easily distinguish a PETase treated plastic sample from a non-treated one and can also provide some information on the degradation velocity. Well-established parameters, which analyze the noise or image quality of a picture, such as variation, standard deviation, SSIM, MSE and PSNR are useful to monitor the alteration of the plastic

CRediT authorship contribution statement

Lena Schnitzler: Conceptualization, Methodology, Software, Formal analysis, Data curation, Writing - original draft, Visualization. Jan Zarzycki: Validation, Formal analysis, Data curation, Visualization, Writing - original draft. Marina Gerhard: Investigation. Srumika Konde: Investigation. Karl-Heinz Rexer: Resources, Investigation. Tobias J. Erb: Resources, Funding acquisition. Uwe G. Maier: Resources, Funding acquisition. Martin Koch: Resources, Funding acquisition. Martin R. Hofmann:

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgments

We would like to thank Marlon Tranelis for designing and 3D printing parts of the LDHM. LS work in this paper was supported by the German Federal Ministry of Education and Research BMBF (Project DigiSeal (16KIS0695)).

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    Current address: Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Str. 10, 35,043 Marburg, Germany.

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