ISO/DIS 24899:2026(en)

ISO TC 61/SC 14/WG 4

Secretariat: DIN

Date: 2025-12-16

Plastics – A method for extraction of microplastics from compost samples

© ISO 2026

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Contents

Foreword v

Introduction vi

1 Scope 1

2 Normative references 1

3 Terms, definitions and abbreviations 1

3.1 Terms and definitions 1

3.2 Abbreviations 2

4 General 3

5 Extraction of microplastics from compost 4

5.1 Principle 4

5.2 Materials, reagents and consumables 5

5.3 Precautions for the laboratory environment, apparatus and materials 5

5.3.1 Operating precautions linked to the laboratory environment 5

5.3.2 Precautions and cleaning protocol for materials 5

5.4 Laboratory Qualification 6

5.4.1 Procedural blank 6

5.4.2 Method blank 6

5.4.3 Verification of microplastics recovery rate 7

5.4.4 Particle stability tests 7

5.5 Deagglomeration 8

5.6 Fenton oxidation 8

5.7 Density separation 8

6 Performance criteria and method verification 10

7 Test report 10

Annex A (informative) Sampling from compost and homogenisation of raw samples 12

A.1 Procedure 12

A.2 Homogeneity test results 13

Annex B (informative) Analysis of extracted microplastics 14

B.1 Sample preparation for analytics 14

B.2 Analytical techniques for the analysis of compost extracts 14

Annex C (informative) Method improvement and validation 15

C.1 Method improvement by an interlaboratory comparison (ILC) 15

C.1.1 General 15

C.1.2 Adaptations in the extraction method 15

C.1.3 ILC results using different analytical techniques (non-standardized) 15

C.2 Stability tests using the final method 15

C.2.1 Particle stability during sonication 15

C.2.2 Particle stability during Fenton oxidation 16

C.3 Recovery tests using the final method 17

C.3.1 Subclause title 17

C.3.2 Subclause title 17

C.3.2.1 Subclause title 17

C.3.2.2 Subclause title 17

Annex D (informative) Recycling of CaCl₂ solution — Procedure 18

Bibliography 19

Foreword

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This document was prepared by Technical Committee ISO/TC 61, Plastics, Subcommittee SC 14 Environmental aspects.

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Introduction

Microplastics are ubiquitous in the environment, occurring also in complex environmental (e.g. soil, sediments) and technical (e.g. compost) matrices. The compost, be it of industrial or home composting origin, is primarily made from organic materials, however contamination can occur if non-compostable plastic waste is not properly separated from the organic materials before composting. Potentially, plastic material fragments into smaller microplastics, while some polymeric materials may undergo complete biodegradation. When compost is used as fertiliser in agricultural soils plastic fragments can enter the environment. Consequently, the presence of plastic of any size not only impacts compost quality but also poses a threat to biota when compost is applied in natural settings.

To analyse microplastics in compost, it is essential to ensure a quantitatively and qualitatively representative extraction. This involves evaluation of plastic particle and polymer stability and high recovery rate, whilst minimising sample contamination. Generally, the extraction of microplastics from compost is poorly comparable between scientific studies due to inconsistent methodologies used and a lack of quality assessments. Therefore, there is a growing need for a standardised method for microplastics extraction from compost for regulatory and standardisation purposes. Furthermore, a potential need for standardised methods for extracting microplastics from compost can be anticipated, as the existing standards, dealing with organic recycling of packaging materials (ISO 17088 or EN 13432), have recently been revised or will undergo a future revision.

This document provides a method for extraction of microplastics from compost, including quality control measures such as recovery and stability tests. Solely the extraction is part of this document. Mass-based or number-based analytical techniques (e.g. according to ISO 16094-2 and ISO 16094-3) commonly used for microplastic quantification can be employed afterwards.

This document covers microplastics from 20 µm to 1 000 µm (in alignment with ISO 16094-2) and with densities below 1,4 g/cm³ (therefore not all types of PVC or PTFE are covered with the current version of this method). Fragments from biodegradable materials will also be extracted with this method.

Plastics – A method for extraction of microplastics from compost samples

This document specifies a laboratory method for the extraction of microplastics from compost matrices originating from industrial or home composting. The method outlines various extraction steps assuring polymer stability, and high recovery rate. This extraction process separates microplastics from the compost matrix that can be further analysed either by number-based or by mass-based techniques.

The method is applicable for microplastics up to 1 mm in size.

The method is applicable for microplastics with densities lower than 1,4 g/cm³.

This document will not specify downstream detection methods for the identification and quantification.

The method in this document has not been validated for microplastic extraction from other matrices, except for composts.

There are no normative references in this document.

For the purposes of this document, the following terms and definitions apply.

ISO and IEC maintain terminology databases for use in standardization at the following addresses:

— ISO Online browsing platform: available at https://www.iso.org/obp

— IEC Electropedia: available at https://www.electropedia.org/

3.1.1

compost

organic soil conditioner obtained by biodegradation of a mixture consisting principally of various vegetable residues, occasionally with other organic material, and having a limited mineral content

[SOURCE: ISO 472:2013, 2.1735]

3.1.2

technical matrix

a matrix specifically designed by controlled combinations of organic materials and additives, often created through human intervention to optimize decomposition and nutrient availability (e.g. compost)

Note 1 to entry: In contrast, natural matrices like soil are complex, dynamic systems formed over long periods through environmental processes, including weathering, organic matter decomposition, and biological activity.

3.1.3

microplastics

solid plastic or synthetic polymer particle insoluble in water with the largest dimension between 1 μm and 5 mm

Note 1 to entry: Microplastics may show various shapes.

Note 2 to entry: This definition encompasses the ISO/TR 21960 definitions of large microplastics and microplastics

Note 3 to entry: The term “microplastics” covers the sum of several individual microplastic particles.

[SOURCE: ISO 16094-2:2025]

3.1.4

sample

small portion of a material or small group of units taken from a larger quantity of material or collection of units and intended to be representative of the whole

[SOURCE: ISO 472:2013, 2.899]

3.1.5

subsample

a smaller, representative portion of a larger sample, used for analysis or testing.

3.1.6

procedural blank

an extraction conducted in absence of microplastics and compost to determine the microplastic contamination from the lab equipment, chemicals and the surroundings.

3.1.7

method blank

an extraction conducted of a compost subsample for the purpose of spike-recovery tests, in absence of spiked microplastics to determine the microplastic background contamination from this specific unspiked compost subsample.

3.1.8

Fenton’s reagent

a solution of hydrogen peroxide (H₂O₂) and an iron catalyst, typically ferrous iron (Fe²⁺), used to generate hydroxyl radicals (•OH) for the oxidation of organic compounds. It is commonly employed in wastewater treatment and environmental remediation to degrade pollutants and is known for its effectiveness in breaking down complex organic materials.

3.1.9

ultrapure water

water that has been purified to an extremely high degree, typically having a resistivity of 18,2 MΩ∙cm and containing minimal levels of contaminants, ions, and organic compounds, making it suitable for sensitive laboratory and industrial applications.

This method is intended for extracting microplastics from compost matrices generated from either industrial or home composting processes.

The method takes into consideration:

a) the extraction of microplastics from 20 µm to 1 000 µm in size;

b) retrieval of microplastics with densities < 1,4 g/cm³, such as polyethylene (PE), polypropylene (PP), polyethylene terephthalate (PET), polycarbonate (PC), polystyrene (PS), polyamide (PA), polymethyl methacrylate (PMMA), polyurethane (PU), as well as retrieval of fragments from biodegradable polymers with densities < 1,4 g/cm³ such as polylactic acid (PLA) and polybutylene adipate terephthalate (PBAT);

c) the heterogeneity of the test matrix (i.e. compost);

d) solely the extraction of the particles. An informative section on sampling is included in Annex A. Several analytical methods were applied in the laboratories during the method’s development. These methods are deemed reliable, and are outlined in Annex B;

e) minimized destructive effects on particle size, number, mass, or chemical composition;

f) reduction of microplastics loss during extraction, due to extensively optimized procedural steps throughout the extraction method;

g) minimization of plastic contamination of sample analysed by excluding, where possible, the use of plastic materials and equipment;

h) control measurements for microplastic contamination, including method and procedural blanks, as well as stability and recovery tests for microplastics and learnings from the interlaboratory comparison as described in Annex C;

NOTE The size and limit of quantification (LOQ) of extracted microplastics is directly correlated with the available detection methods for their identification and quantification and can therefore not be normative in this document. The current version of this document is considered robust for the size range of 20 µm to 1 000 µm. The LOQ can be as low as one particle with 20 µm in size per gram of compost when the entire extract is analyzed, provided that no contamination of the target analyte is detected in the procedural blank. However, if only one particle is identified in a subsample of the extract (for instance, one-fifth of the total extract), the LOQ for that single particle must be multiplied by five in this context. Additionally, if any contamination of the target analyte is present in the procedural blank, the number or mass of detected microplastics must exceed ten times the level found in the blank to be statistically significant. Therefore, minimizing or eliminating contamination in the procedural blank is crucial for achieving a lower LOQ. Nonetheless, future revisions of this document could focus on broadening the size spectrum and incorporate methods for extracting and analysing smaller microplastics, provided that the current analytical challenges we are facing are overcome.

The extraction method for microplastics from the compost involves several sequential steps. It starts with the deagglomeration of the compost particles and the oxidation of the organic material using Fenton oxidation. Microplastics are collected following density separation.

The extraction procedure has been optimized by an interlaboratory comparison, as referenced in Annex C, which ensures a higher level of quality and reliability of the standard. If any steps of the extraction are omitted, the performance criteria provided in Clauses 4 and 6 of this document become invalid, since the same level of quality (extraction efficiency and matrix removal efficiency) in the extraction cannot be guaranteed.

A detailed overview of steps, duration of the extraction of microplastics from compost, as well as reagents and materials needed, are given in Table 1.

Table 1 — Detailed overview of steps and duration of the extraction of microplastics from the compost

NOTE The steps of homogenisation, representative sampling, and subsequent detection of particles (identification and quantification of microplastics) are not part of this document. For further information on these two steps please look to Annexes A and B.

The water and all chemicals listed in Table 1 need to be checked for the presence of microplastics by applying the procedural blank approach in 5.4.1. The limit of quantification is defined by the background contamination. The particle number or mass in the analysed compost sample must be 10 times higher than the background contamination determined by the procedural blank measurements. In case of high procedural blank contamination, microplastics in the chemical solutions should be removed by filtration (for example, through a cellulose filter, metal filter or another non-polymeric membrane with a pore size of max. 20 µm) to achieve a lower limit of quantification. In case of targeted analysis, no equipment made of the target polymer shall be used.

The space dedicated to preparing and filtering samples should be free from polymer coatings or materials. Regular cleaning of the laboratory environment is mandatory (e.g. hood and lab bench). To do this, ethanol, detergent, particle-free water and suitable polymer free wipes may be used. In particular, operators shall:

— check potential sample contamination by microplastics and chemicals which may cause spectral interference with microplastics via the procedural blank measurements;

— use new gloves to avoid or minimize the unwanted release of microplastics from the laboratory safety equipment;

— wash their hands prior to starting the manipulations with samples, especially after washing the outsides of containers and when entering rooms dedicated to handling samples;

— wear a cotton lab coat or, if necessary, a clean anti-static lab coat (e.g. characterised regarding on the materials/polymer in order to exclude the type of polymer from the sample results report);

— not wear face masks made of synthetic polymers; not wear clothing made of synthetic fibres (fleece jackets for example), body cleansing products or cosmetics are likely to release microplastics in the laboratory environment (e.g. nail varnish, foundations, exfoliating products);

The laboratory personnel should protect the sample from all contamination coming from the working environment especially when transporting the sample between the preparation workstation and the analysis workstation (e.g., by covering the sample with aluminium foil or placing it in a suited container), and during this final analysis stage.

Small equipment that can release microplastics such as those analysed (PP, PC, PA, etc.) should not be used.

All items (glassware, metal etc.) getting in contact with the samples, including containers for sampling, shall be subject to special attention and shall be cleaned thoroughly, by applying the following protocol or other suitable cleaning procedures:

Immerse the glass items in a suitable detergent solution, with a sufficient contact time. Then rinse the items with a suitable product (e.g. 50 % ethanol or a neutralising agent) and complete rinsing with particle-free water of known quality. Leave the equipment to air dry. Do not wipe it dry.

As an alternative, glass items can be filled with particles free water of known quality and surfactant and put in an ultrasonic bath for 5 min. The water is removed, and the entire procedure is repeated two times more.

Containers, glassware and utensils may be calcined at 480 °C for 2 h or 450 °C for 6 h.

While working in a clean room is not strictly mandatory for the extraction outlined in this document, it is however important to take all measures to avoid contamination of the samples. In addition, it is essential to analyse procedural blanks to determine the microplastic background from the surrounding area and laboratory equipment. While plastic equipment should generally be avoided, it may be used in exceptional cases for targeted analysis, provided that the plastic material can be distinguished from the target analyte.

To create a procedural blank sample, the extraction steps outlined in 5.5 to 5.7 shall be simulated in the absence of compost. Step 5.5 will yield a 100 ml sample of ultrapure water, which is continuously stirred for 3,5 h. After the set period, the stirred ultrapure water is mixed with additional 200 ml of ultrapure water and then 375 g CaCl₂ dihydrate (99 %) is added, generating a 55 wt.-% salt solution. The salt solution is transferred into a separating funnel, leaving the mixture to settle for 18 h. When this period has passed, 25 ml of the 0,036 M Fe²⁺ solution will be filtered through a 20 µm filter and it will be rinsed with 50 ml of ultrapure water. The content of the filtration flask shall be discarded before the next filtration. Following, the main proportion of the salt solution in the separating funnel is discarded and only the last 100 ml of the separating funnel content is filtered through the same 20 µm filter. The separating funnel is then rinsed with 50 ml water and 100 ml ethanol and the rinsing solution is also filtered through the same 20 µm filter. The filter is again rinsed with 50 ml of ultrapure water and the filtration flask content is discarded. Finally, 50 ml of H₂O₂ (30 wt.-%, liquid) are filtered through the same 20 µm filter. Subsequently, a spray bottle filled with 100 ml of ultrapure water is used to rinse any remaining particles from the metal filter into a 250 ml bottle. It is essential to analyse the entire content of the 250 ml bottle. The count/mass of microplastics present corresponds to the background of microplastic count/mass in the final 100 ml analytical sample after the 20 µm filtration step. The background of microplastic count/mass shall be recalculated if a subsample is taken for compost analysis. The LOQ of the extraction and analysis is directly correlated with the analytical technique used for quantification after extraction (see Note in Clause 4). Additionally, if any contamination of the target analyte is present in the procedural blank, the number or mass of detected microplastics must exceed ten times the level found in the blank to be statistically significant. Therefore, minimizing or eliminating contamination in the procedural blank is crucial for achieving a lower LOQ.

For spike-recovery tests, it is crucial to perform method blank measurements. The method blank comprises a 1 g compost sample that does not have any intentionally spiked microplastics or plastic test items. This 1 g compost subsample should be sourced from the same compost sample of which later another 1 g subsample will be spiked with the target analyte. To assess microplastic contamination in this method blank, the complete extraction process outlined in sections 5.5 to 5.7 must be conducted, and the analysis should employ the same analytical technique that will be used for the spiked samples. The method blank assessment is only possible for the spike-recovery tests and is identical to the sample that needs to be analysed for microplastic content.

Verification of the successful implementation of the extraction method may be checked using at least one microplastic material of a known type (among the ten main ones).

The use of commercial polymer/microplastic standards (e.g. PS beads or standards used for flow cytometry) or a reference prepared by the laboratory (by friction, grinding, etc.) is possible.

Number characterization of the prepared spike sample may take place using a different analytical technique (e.g. flow cytometry, optical counting, Micro-FTIR, Micro-Raman, SEM) from the one used for quantification after extraction.

Spike at least three samples of a compost method blank (5.4.2) with a known mass or number of microplastics of different types and sizes to cover the field of application claimed by the method. The complete extraction process outlined in sections 5.5 to 5.7 must then be conducted, and the analysis should employ the same analytical technique that will be used for the method blank.

The recovery rate is calculated by subtracting the mass or number of microplastic found in the method blank from the mass or number found in the spiked samples. As an additional reference, the same number or mass of microplastics must be spiked into 100 ml of water and analysed using the same analytical technique employed for the method blank and the spiked sample. This approach allows for the differentiation of analytical deviations from those related to the extraction method.

It should be ensured that the recovery rate is >60 % for particles < 100 µm and >80 % for particles > 100 µm. If not, the (analytical) method should be optimised. For values above 100 % the laboratory will have to perform a cause analysis.

Several types of microplastics and fragments from biodegradable polymers were tested during method development regarding their stability towards the specific sonication and Fenton oxidation conditions applied in this standard (results shown in Annex C). If any other type of microplastic or more complex materials, such as multicomponent materials containing fillers and not predominantly composed of pure polymers, are to be investigated using this standard, it is essential to conduct stability tests on the target analyte. This ensures that neither the particle size distribution nor the mass is altered by more than 10 %. If the stability test results indicate a change of 10 % to 30 %, the impact on the reported results must be discussed in the test report. If the alteration exceeds 30 %, the target analyte cannot be extracted using this protocol.

To test the stability of the microplastic type of interest under the conditions of the described extraction procedure (5.5 to 5.7), a suspension of the microplastics of interest in ultrapure water (in the absence of any compost matrix), is subjected to the different steps of the process as foreseen in this document. The particle size distribution or mass of each particle (polymer) type are analysed before and after applying the extraction procedure and the results are compared.

It is also possible to test the particle stability in single extraction steps only (e.g. 5.6 Fenton oxidation) to identify the most critical step for the stability of the microplastic type of interest.

The minimum requirement for analysis is to compare the particle size distribution (e.g. laser diffraction: Fraunhofer evaluation for particles > 50 µm, MIE evaluation for particles 1 µm to50 µm) or mass of the microplastic type before and after carrying out the whole procedure (or steps of it). Additional analyses can be conducted based on the specific microplastic type and research question. Examples of additional analyses include gel permeation chromatography for molar mass distribution (applicable only to non-crosslinked types of microplastics), (ATR-FT)IR and/or XPS for surface chemistry, optical microscopy or SEM for surface texture and shape, and DSC for crystallinity.

After homogenization and subsampling, any agglomerates of microplastic and compost particles are broken down and dispersed using ultrasound. To achieve this, 1 g of compost subsample and 40 ml ultrapure water are added to a beaker (e.g. 100 ml) and ultrasonicated with a delivered power of 15,0 W to 20,0 W for 5 min (corresponding to 4 500 J to 6 000 J) to effectively break up the agglomerates. The device used for ultrasonication shall have its delivered power measured using the calorimetric method described by Taurozzi, Hackley and Wiesner (2012). Once the ultrasonication is complete, the ultrasonication tip is removed and rinsed with 10 ml of ultrapure water directly into the same beaker containing the 40 ml compost dispersion. The resulting dispersion is transferred to a beaker (e.g. 500 ml to 600 ml, ⌀ 8 cm) and stirred using an overhead stirrer with a ceramic or glass rod. The 100 ml beaker is rinsed with 50 ml ultrapure water. Rinsing steps throughout the method reduce microplastics loss due to attachment to the labware used.

NOTE The ultrasonication was tested using the Branson digital probe Sonifier SFX 550 with a micro-tip (6,4 mm, 1/4”, conical)^[1]. To achieve the optimal delivered energy, the test matrix was ultrasonicated for 5 min at 40 % intensity.

The majority of organic components of the compost matrix will be removed using a Fenton oxidation procedure at room temperature. For safety reasons, this oxidation step must be conducted in a fume hood. For each extraction, a solution of 0,036 M Fe²⁺ (obtained by dissolving 10 g FeSO₄ heptahydrate in 1 l of ultrapure water) needs to be freshly prepared and the pH will be tested and adjusted to 3 to 4 with a 1 M HCl (liquid) or a 1 M H₂SO₄ (liquid). The oxidation is performed in five steps. In each step, Fenton reagents, 10 ml H₂O₂ (30 wt.-%, liquid) and 5 ml 0,036 M Fe²⁺ solution, are mixed directly into the compost sample beaker while stirring at 250 rpm. The next portion of Fenton reagent is added at every 30 min of continuous stirring. Following the last addition (= fifth addition) of the Fenton’s reagent, the compost/microplastics dispersion is stirred continuously for 60 min. Then the dispersion is left to sediment in the beaker for 60 min. Only the supernatant of the dispersion is carefully filtered through a 20 µm metal filter to remove the Fenton reagent. Approximately 10 g of the dispersion containing the sediment is retained in the beaker. This precaution ensures that microplastics do not become trapped in a filter cake, which could lead to recovery challenges in the subsequent step 5.7 density separation. It is essential that the particles on the filter remain moist and are processed without delay in the following step 5.7 density separation.

NOTE 1 The Fenton’s reagent used in this procedure does not impact the sizes and molar masses of the microplastic. The stability of PBAT, a PLA-based blend, LDPE, PET was assessed during the method’s development (Annex C). This type of stability test should be conducted once in each laboratory unless literature data for the target analyte is available.

NOTE 2 Fenton oxidation is a chemical reaction and requires safety precautions: perform oxidations in a fume hood, monitor reaction intensity and temperature closely, and ensure the temperature does not exceed 50 °C.

NOTE 3 Fenton’s reagent is a solution containing iron ions, which contribute to the formation of an orange residue (Fenton residue) during Fenton oxidation. The residue, which was not removed during the 20 µm sieving step, is removed in the following step (5.7 Density separation).

It is imperative to use analytical-grade CaCl₂ dihydrate (99 %) rather than CaCl₂ anhydrous, to achieve the desired high density and viscosity of the salt solution, as well as to ensure its specified purity. The salt solution should be prepared at room temperature in a separate beaker by stirring 375 g of CaCl₂ dihydrate (99 %) in 300 ml of ultrapure water. This results in a 5,24 M Ca²⁺ or 55 wt.-% CaCl₂ dihydrate (99 %) salt solution, with a density of 1,4 g/cm³.

To ensure complete transfer of the microplastics, the particles on the 20 µm metal filter are rinsed back into the beaker containing the sediment by using 100 ml of 55 wt.-% CaCl₂ dihydrate solution. To effectively carry out the density separation, a separating funnel (at least 1 l capacity, borosilicate) with a closing valve at its end is filled with prepared 100 ml CaCl₂ dihydrate solution. For easier removal of the high-density fraction from the separating funnel, a mark of 100 ml is painted with a permanent marker on the funnel’s surface. A smaller glass funnel is placed on top of the separating funnel. The compost and microplastic particles in the beaker are transferred into the separating funnel and the beaker is rinsed with 200 ml of CaCl₂ dihydrate solution, dispensed directly from a spray bottle into the funnel. It is imperative that the high-density salt solution is added first into the funnel, followed by the compost/microplastic particles, to improve the microplastics recovery. The upper part of the funnel is closed with a glass lid to prevent contamination. The separation funnel with 400 ml compost/microplastics dispersion is first swirled horizontally in circles for 1 min to 2 min so that its content begins to rotate and forms a vortex. It is crucial that the funnel in not shaken in the usual way to prevent contamination of the lid. The dispersion is then left to sediment. After 18 h, the salt solution containing the high-density fraction and Fenton residues is removed by slowly opening and closing the valve at the bottom side of the separating funnel, in intervals of 30 min for a period of 6 h until 100 ml of the liquid is left in the funnel. Another 200 ml salt solution is then added to the separation funnel, and the funnel is horizontally swirled in circles as described above and left to sediment for the second time. After 18 h, 200 ml of the salt solution is removed again by opening the valve at the bottom of the separating funnel. The density of the final salt solution (the removed portion containing the high-density fraction, such as Fenton residue) should be checked by weighing 10 ml in a graduated cylinder on a balance. The remaining 100 ml of the supernatant with floating microplastics is retained in a 250 ml glass bottle. The separating funnel is rinsed first with 50 ml ultrapure water followed by 100 ml ethanol into the same 250 ml glass bottle. To remove any remaining solid residues which might disturb the analysis, the sample is filtered through a metal filter with a pore size of 20 µm. Next, a spray bottle containing 100 ml of ultrapure water is used to rinse the particles from the filter back into the 250 ml sample container. The entire content of the spray bottle must be utilized, even if this requires opening the bottle at the end and pouring out any remaining liquid instead of spraying it. This constitutes a final sample.

NOTE 1 To promote environmental sustainability and reduce expenses, the CaCl₂ solution can be conveniently reused (refer to Annex D for more information).

NOTE 2 Other salt solutions were initially tested for density separation, but the best reproducibility was achieved using a CaCl₂ solution. Sodium polytungstate (SPT) represents the only alternative for extracting both microplastics and biodegradable fragments with densities greater than 1,4 g/cm³. However, since this method has not been validated with SPT, the performance criteria outlined in Clause 6 of this document are no longer applicable, as the same quality of extraction efficiency and matrix removal efficiency currently cannot be assured. Consequently, this adaptation of the method must undergo validation through spike-recovery and particle stability tests, as well as blank measurements. While the use of sodium iodide (NaI) has the potential to extract microplastics with higher density, it is a more expensive option and often results in the formation of elemental iodine. This can cause a yellowish discoloration of the solution, as well as laboratory equipment and microplastics. Additionally, changes in the density of the salt solution cannot be excluded. Similarly, although zinc chloride (ZnCl₂) is more affordable, it was found to be less efficient in the density separation step. The recoveries of microplastics were lower compared to the CaCl₂ solution due to its interaction with the Fenton residue. Furthermore, the ZnCl₂/Fenton mixture has the potential for precipitate formation, which can entrap microplastics and cause also false negative or positive results for spike recovery when applied to samples with smaller microplastics. Salts that produce basic solutions, such as potassium bromide (KBr), are not suitable for extracting biodegradable fragments, as these materials are likely to degrade under alkaline conditions.

For the extraction to be considered valid, the following performance criteria shall be met:

a) All steps 5.5 to 5.7 of the extraction method and at least one blank measurement (5.4.1 or 5.4.2) in triplicate shall be carried out.

b) Spike recovery tests as outlined in 5.4.3 shall be conducted at least once per laboratory and a recovery of > 60 % shall be achieved for particles < 100 µm and > 80 % for particles > 100 µm. For untargeted analysis, if a microplastic reference material is available and suitable in size and composition, it shall be used for spike-recovery tests (5.4.3), and results shall be reported. For targeted analysis, a microplastic test or reference material with the same chemical composition as the target analyte shall be used.

c) The delivered power of the ultrasonication device shall be determined at least once per laboratory using the calorimetric method described by Taurozzi, Hackley and Wiesner (2012) and shall be in the range of 15,0 W to 20,0 W.

d) Particle stability in the Fenton’s reagent shall be confirmed by stability tests as outlined in 5.4.4 and Annex C, particularly when data is not available from the literature.

e) The final density of the salt solution in the separating funnel shall be determined and shall be at least 0,05 g/cm³ higher than the density of the target analyte.

f) Plastic contamination of the sample shall be avoided throughout the procedure (laboratory equipment and clothing). Data of the procedural and/or method blank shall be reported and discussed in context with the results.

The test report shall provide all relevant information, including:

a) the test method used, together with a reference to this document, i.e. ISO/DIS 24899:2026;

b) all information required to identify the sample (available from composter):

— Origin of compost (plant operating company, plant location, type of plant, size of plant)

— Date of production

— Storing conditions (time, temperature, humidity)

— Rotting time

— Finishing procedure (e.g. sieving, sieve size)

c) date the laboratory received the sample and date of sample analysis;

d) list of plastic equipment utilized during the extraction process, including its chemical identity, if applicable

e) delivered power during sonication

f) final density of the salt solution in the separating funnel and delivered power of the ultrasonication device,

g) spike recovery test results using microplastic reference materials if available or other microplastics of interest; reporting in counts or mass per gram dry compost (5.4.3 and Annex C),

h) information concerning the type, number/mass and smallest size of the microplastic reference material. This will allow for determining the smallest size of microplastics (in µm) that can be extracted and detected. The detection of microplastics depends on the chosen detection methods, the density of the target analyte and pre-filters used for the analysis,

i) results of blank measurements: 1) procedural blank (extraction in absence of compost), 2) method blank (compost without spiked microplastics for comparison to the spiked sample); reporting in numbers or mass per gram dry compost,

j) any event or operation not described in this document that can affect the result shall be reported.

1. 
   (informative)
   
   Sampling from compost and homogenisation of raw samples
   1. Procedure

The sampling procedure for compost and subsequent homogenization are not normative. However, a recommended sampling method that has demonstrated suitability is presented below.

Prior to the extraction procedure, collect compost from industrial composting plants in metal buckets (min. 5 l). The initial sample should be manually shaken in all directions and divided into subsamples of 1 l each, placed in glass bottles with a minimum volume of 2 l. If possible, avoid the use of a plastic cap or report the polymer type used. When shaking one of the glass bottles, it is crucial to periodically rotate the bottle instead of shaking it repetitively in a unidirectional manner to avoid uneven mixing, which can result in inaccurate or inconsistent results. One of these subsamples is then taken and shaken again, periodically rotating the bottle during the shaking process. The 1 l subsample is subsequently divided into 100 ml glass bottles. Five of these bottles are manually shaken, combined in a larger glass container, and shaken once more. For further processing, a 100 ml subsample is obtained.

The 100 ml subsample undergoes freeze-drying and is then transferred into a sieving device fitted with a 1 mm sieve. To ensure the break-up of any agglomerates in the compost, the sieving device is supplemented with 30 stainless-steel spheres (⌀ 5 mm, 240 min, amplitude of vibration 1,5 mm). The resulting homogenized subsample exhibits particle sizes < 1 mm, which is the maximum size of the microplastics targeted in the extraction process. Special attention should be given to the visual aspects of the compost. The organic residue should be indistinguishable to the naked eye at a distance of 500 mm. The visible assessment of the compost matrix (independent of microplastic presence) should be documented through photography (adopted from ISO 17088:2021).

Before obtaining 1 g of compost for the extraction, the 100 ml homogenized subsample is thoroughly mixed manually or in a shaker mixer. In the case of the shaker mixer, the compost is mixed using a combination of rotating, translational and inverting motions.

A grinding approach can be considered to achieve homogeneous samples only if the chosen analytical technique for microplastic quantification is a mass-based method that does not differentiate between microplastic numbers and sizes.

NOTE 1 If a sieving machine is not available, manual grinding and sieving of the selected 100 ml sample through a 1 mm stainless-steel metal sieve could be done as well. The griding and sieving should be done until no breakable agglomerates of compost remain in the sample, this should be visually checked by the operator.

NOTE 2 Homogenization of the test matrix was assessed using 600 g dry compost in a 1 l glass container. The container was placed in a 3D shaker mixer (TURBULASYSTEM SCHATZ T2F^[2]) for 30 min. Through visual inspection (adopted from ISO 17088:2021) and duplicate measurement of extracted samples containing microplastics, the homogeneity of the compost was confirmed. As an alternative to the shaker mixer, manual mixing, as described in the main text, can also be utilized. To conduct homogeneity testing of the test matrix, follow these steps: Add a known quantity of fluorescent microplastics (> 125 µm) to 100 g of dry compost. Thoroughly mix the mixture manually and then take out a 1 g subsample. Finally, manually count the fluorescent microplastics using a fluorescence microscope.

• 1. Homogeneity test results

The homogeneity of 1 g compost subsamples was checked via thermogravimetric analysis (TGA) by comparing their decomposition curves. For this purpose, 100 g of dry compost < 1 mm was mixed a) with the 3D shaker mixer and b) by manual mixing as described above before each 1 g subsample was taken out. The whole 1 g subsample was measured from 24 °C to 550 °C. Only the first subsample that was taken out after the 3D shaker approach showed a higher deviation of >10 % from the other subsamples (Figure A.1).

To test the homogeneity of particle numbers in compost subsamples, 20 g of autoclaved and sieved (< 1 mm) compost were mixed manually with a small amount of fluorescent PBAT particles (100 µm to 300 µm). 1 g compost subsamples were analysed using a fluorescence microscope. For the procedure, 1 g of the mixture was weighed, then distributed on a petri dish, and all the fluorescent particles present on that filter counted. Results from this homogeneity test showed an average particle count per gram of compost of 34,85 particles with a standard deviation of 12,25. With a median count of 35, the 5 % percentile of 16,85 and the 95 % percentile of 53,55.

These results demonstrate that homogeneous subsampling from freeze-dried compost < 1 mm is possible, however, a separate standard is needed to elaborate on this approach.

Figure A.1 — TGA curves from 1 g compost subsamples taken after using either a 3D shaker or manual shaking.

1. 
   (informative)
   
   Analysis of extracted microplastics
   1. Sample preparation for analytics

If necessary, the supernatant can be subsequently vacuum filtered through a sieve or a cascade of sieves to obtain subsamples of different size bins of microplastic particles. This step also enables the removal of solid residues smaller than 20 µm, if using the appropriate pore size of the sieve. The microplastics can then be quantified by mass-based spectrometric techniques, such as py-GC/MS or TED-GC/MS, and/or using various microscopy techniques, such as Raman microscopy, fluorescence microscopy, (FT)IR (Fourier Transform Infrared) microscopy or LDIR (Laser Direct Infrared) microscopy.

• 1. Analytical techniques for the analysis of compost extracts

It is highly recommended to analyse the extracts according to ISO 16094-2 or ISO/DIS 16094-3 with one of the analytical techniques described therein.

1. 
   (informative)
   
   Method improvement and validation
   1. Method improvement by an interlaboratory comparison (ILC)
      1. General

During the ILC the participants gave detailed feedback on the method, difficulties they found and how to solve the issues they encountered. This led to a series of changes in the used equipment and the methodology of this standard. Here the comments/changes are presented in the internal chronology of the methodology.

• • 1. Adaptations in the extraction method

The use of a glass encapsulated stirring bar was deemed unsuitable, given that these are prone to breaking and abrasion, potentially compromising the sample and releasing glass shards that add particles to the posterior analysis. Favouring and leaving as the only option the use of an overhead stirring device with a glass or ceramic rod. The different laboratories noticed that the used Fenton reagent proportions of hydrogen peroxide and iron solution could lead to an elevated iron oxide particle formation and deposition on the polymer surface. This problem is shown during the spectroscopy analysis techniques, which then cannot identify correctly the polymer particle. This detected problem led to the reduction and the adjustment of the use of iron solution in the organic matter treatment phase of the method, leading to the currently stated concentrations. Finally, several participants reported the precipitation of CaCl₂ or the formation of a foam during the addition of CaCl₂ to the compost and microplastic dispersion after Fenton oxidation. It is believed that the temperature, unreacted hydrogen peroxide and a component of the compost promoted the foam formation. The reports by the ILC partners led to the complete change of the density separation preparation step, resulting in the current state of the methodology that includes an intermediate filtration at 20 µm to avoid dissolving the CaCl₂ directly in the dispersion. This will also allow a better control of the CaCl₂ concentration during the density separation, avoiding a possible crystallization of the salt.

• • 1. ILC results using different analytical techniques (non-standardized)

New references were sent out to the ILC partners, results will be included as soon as the results on the reference samples have been received.

• 1. Stability tests using the final method
     1. Particle stability during sonication

The particle stability during sonication according to subclause 5.4.4 was assessed with the final protocol, which was improved by the input of partners from the interlaboratory comparison.

The particle size distributions of the polymer particles were determined via Fraunhofer light scattering before and after sonication using a Mastersizer 3 000 (MV Hydro unit) with a measuring accuracy of 0,6 %. The sample volume was determined by adjusting the light shading range to between 5 % and 16 %. The surfactant Nekanil 910 was added to the sample. As shown in Figure C.1, the particle size distribution of the investigated powers was altered by << 10 %.

Figure C.1 — Measured particle size distributions of PBAT, LDPE, PET before and after sonication.

• • 1. Particle stability during Fenton oxidation

The particle stability during Fenton oxidation according to subclause 5.4.4 was assessed with the final protocol, which was improved by the input of partners from the interlaboratory comparison.

The particle size distributions of the polymer particles were determined via Fraunhofer light scattering before and after oxidation using a Mastersizer 3 000 (MV Hydro unit) with a measuring accuracy of 0,6 %. The sample volume was determined by adjusting the light shading range to between 5 % and 16 %. The surfactant Nekanil 910 was added to the sample. The total mass (approx. 200 mg) of the polymer powders before and after oxidation was determined by weighing. As shown in Figure C.2, the particle size distribution of the investigated powders was altered by << 10 % and the mass recovery was 99,3 wt.-% for a PLA-based blend, 98,7 wt.-% for PBAT, 99,2 wt.-% for PET, and 99,6 wt.-% for LDPE.

Figure C.2 — Measured particle size distributions of a PLA-based blend, PBAT, PET, LDPE before and after Fenton oxidation.

Without compost, test conditions are more severe than with compost (Figure C.3). Therefore, if a polymer passes the stability test without compost, it will also be stable in the presence of compost. In all cases, temperatures remain below 40 °C.

Figure C.3 — Temperature evolution monitored during stability tests in absence of compost (blue curve) and in presence of compost (orange and grey curves).

• 1. Recovery tests using the final method

Information will follow.

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1. 
   (informative)
   
   Recycling of CaCl₂ solution — Procedure

Following the extraction of microplastics, the CaCl₂ solution can be recycled. To do this, the liquid collected from the separating funnel after density separation is filtered through a 1 µm pore size glass fibre filter (⌀47 mm). When using the recycled CaCl₂ solution in the next extraction, the density should be checked in a gradual cylinder by weighing 10 ml of the solution. If the density is lower than 1,4 g/cm³, it should be adjusted by gradually adding CaCl₂ dihydrate.

NOTE The filtrate can have various colours, ranging from translucent to orange. The colour does not affect the performance of the density separation.

Bibliography

EN 13432, Packaging — Requirements for packaging recoverable through composting and biodegradation — Test scheme and evaluation criteria for the final acceptance of packaging

ISO 472, Plastics — Vocabulary

ISO 3310‑1, Test sieves — Technical requirements and testing — Part 1: Test sieves of metal wire cloth

ISO 5430, Plastics — Ecotoxicity testing scheme for soluble decomposition intermediates from biodegradable plastic materials and products used in the marine environment — Test methods and requirements

ISO 16094‑2, Water quality — Analysis of microplastic in water — Part 2: Vibrational spectroscopy methods for waters with low content of suspended solids including drinking water

ISO/DIS 16094‑3, Water quality — Analysis of microplastic in water — Part 3: Thermo-analytical methods for waters with low content of suspended solids including drinking water

ISO 16929, Plastics — Determination of the degree of disintegration of plastic materials under defined composting conditions in a pilot-scale test

ISO 17088:2021, Plastics — Organic recycling — Specifications for compostable plastics

ISO/TR 21960, Plastics — Environmental aspects — State of knowledge and methodologies

ISO 24187, Principles for the analysis of microplastics present in the environment

Braun, U., Altmann, K., Herper, D., Knefel, M., Bednarz, M., & Bannick, C.G., Smart filters for the analysis of microplastic in beverages filled in plastic bottles. Food additives & contaminants. Part A, Chemistry, analysis, control, exposure & risk assessment 2021, 38, (4), pp. 691-700.

Brinton, W. F., Characterization of man-made foreign matter and its presence in multiple size fractions from mixed waste composting. Compost Science & Utilization 2005, 13, (4), pp. 274–280

Pfohl, P.; Roth, C.; Meyer, L.; Heinemeyer, U.; Gruendling, T.; Lang, C.; Nestle, N.; Hofmann, T.; Wohlleben, W.; Jessl, S., Microplastic extraction protocols can impact the polymer structure. Microplastics and Nanoplastics 2021, 1, 13 pages

Traurozzi J.S., Hackley V.A., Wiesner M.R. Preparation of Nanoparticle Dispersions from Powdered Material Using Ultrasonic Disruption, 2012, NIST Special Publication 1200-2, Version 1.1, 14 pages

1. Sonifier SFX 550^® is a trademark of Branson Ultrasonics Corporation, USA. This information is given for the convenience of users of this document and does not constitute an endorsement by ISO of the product named. ↑

2. TURBULASYSTEM SCHATZ T2F^® is a trademark of WAB Willy A. Bachofen AG, Germany. This information is given for the convenience of users of this document and does not constitute an endorsement by ISO of the product named. ↑