A validated Nile red fluorescence method following KOH/H₂O₂ digestion for microplastic detection in marine molluscs

Graphical abstract

1How to digest marine molluscs (biota)?

Fig. 1 digestion protocol — **Fig. 1.** Protocol for processing and digestion of biological samples (e.g., *M. galloprovincialis* and *R. venosa*) using a two-step digestion method. Open the interactive version ↗

2Why is an additional (second) digestion step with 30% H₂O₂ needed?

Fig. 2 orange versus green filters — **Fig. 2.** First line (A–D): orange filters – *M. galloprovincialis* specimens were digested following only stage 1 of the digestion protocol (10% KOH at 60 °C for 24 h) and filtered on Whatman GF/D filters. A strong orange fluorescent signal is visible, which hinders identification. Second line: green filters – *M. galloprovincialis* specimens were digested following stage 1 and stage 2 of the digestion protocol (10% KOH and 30% H₂O₂ at 60 °C) and filtered on Whatman GF/D filters. The orange fluorescent background changes to green, allowing easier identification of the particles.

3Validation of the digestion method

How fluorescent are virgin synthetic polymers after staining with Nile red dye?

Drag the handle (or use the arrow keys) to compare Fig. 3A and Fig. 3B.

Fig. 3A / 3B. Virgin synthetic polymers after staining with Nile red.

How does the fluorescence of synthetic polymers change after the digestion method?

**Fig. 4.** Synthetic polymers and their fluorescence before and after applying the digestion method. Rows indicate imaging conditions: brightfield white light pre-digestion (top), fluorescence imaging (λ = 470 nm excitation; yellow bandpass filter) pre-digestion (middle), and fluorescence imaging post-digestion (bottom).

How fluorescent are virgin natural materials after staining with Nile red dye?

Drag the handle to compare Fig. 5a and Fig. 5b.

Fig. 5a / 5b. Virgin natural materials after staining with Nile red.

How does the fluorescence of natural materials change after the digestion method?

**Fig. 6.** Natural fibres and their fluorescence before and after applying the digestion method. Rows indicate imaging conditions: brightfield white light pre-digestion (top), fluorescence imaging (λ = 470 nm excitation; yellow bandpass filter) pre-digestion (middle), and fluorescence imaging post-digestion (bottom).

How does the fluorescence intensity change after the digestion process?

Before digestionAfter digestionHover or tap a bar for values · error bars = SD (n = 3)

Bar chart 1. Fluorescence intensity (normalized intensity – IntDen/mg) before and after digestion protocol for all 12 materials (6 synthetic polymers and 6 natural fibres). SD is based on 3 replicates. Error bars represent standard deviation from 3 replicates.

< 30% loss30–60% loss> 60% loss

Bar chart 2. Percent change in fluorescence intensity after digestion protocol, color-coded by magnitude (green: <30% loss, orange: 30–60% loss, red: >60% loss).

How does the colour of the materials change after the digestion process?

**Fig. 7.** Fluorescence colour changes represented as hex colour codes before and after digestion. Synthetic polymers maintain orange/yellow tones, while natural fibres lose fluorescence or shift to darker colours.

How much of the material has degraded after the digestion process?

Synthetic polymersNatural materials

Bar chart 3. Material degradation during digestion protocol, showing net material loss after accounting for transfer losses.

Does the digestion protocol affect the chemical integrity of the synthetic polymers?

**Fig. 8.** ATR-FTIR spectra for all polymers before and after applying the digestion protocol, with the fingerprint region (600–1800 cm⁻¹) highlighted in grey.

Hit Quality Index (HQI) values ranged from 89.71% to 99.31%. PE (90 µm) and PET (250 µm) demonstrated excellent spectral agreement (HQI ≥ 95%), while PP (90 µm), PA-66 (100 µm), and PE (35 µm) showed good agreement (HQI: 91.72% – 93.05%). PMMA (250 µm) had slightly lower values (89.71% HQI), however pre- and post-degradation spectra showed good similarity. Furthermore, staining the MP particles with NRD did not appear to interfere with ATR-FTIR identification under the conditions used in this study.

4Hardware components for capturing the filters

**Fig. 9.** Hardware components of a semi-automatic device for imaging fluorescent microplastic particles.

5Creating a panorama with the free AutoStitch software

**Fig. 10.** Creating a panorama with the free AutoStitch software.

6Software for microplastic detection – Microplastic Annotation Package by Park et al., 2022

Guide video

7Testing the method using natural samples of Mytilus galloprovincialis and Rapana venosa from the Black Sea

M. galloprovincialis1.3median MPs g⁻¹ WW · n = 27

R. venosa0median MPs g⁻¹ WW · n = 40

Maximum14.9MPs g⁻¹ WW (R. venosa)

Stations7S1–S7

M. galloprovincialisR. venosa◆ mean · line = median · box = Q1–Q3 · whiskers = min–max

Boxplots. Microplastics per gram wet weight (MPs g⁻¹ WW). M. galloprovincialis (median 1.3; Q1–Q3 0–3.2; mean 2.2; min 0; max 9.9; n = 27) and R. venosa (median 0; Q1–Q3 0–0.2; mean 1.4; min 0; max 14.9; n = 40).

Where were the specimens collected?

Tap a station marker for coordinates and results. Switch between satellite and satellite + labels with the layer control.

Loading map…

Mean values ± standard deviation for tissue weight and shell size. MPs g⁻¹ WW and MPs ind.⁻¹ presented as medians (Q1–Q3) for the seven stations by species.

What does an actual sample of Mytilus galloprovincialis look like?

Drag the handle to compare Fig. 11a and Fig. 11b.

Fig. 11a / 11b. An actual sample of Mytilus galloprovincialis.

What does an actual sediment sample from the seafloor look like after microplastics have been extracted by density separation using K₂CO₃ (1.56 g cm⁻³)?

Drag the handle to compare Fig. 12a and Fig. 12b.

Fig. 12a / 12b. Sediment sample from the seafloor after density separation with K₂CO₃ (1.56 g cm⁻³).

Shells with cavities contain air, which makes it very difficult to extract microplastics completely (without sediment residues) based on density. Some microplastics are difficult to see with the naked eye without Nile red staining.

How much does sample analysis cost?

Cost category	Component	Price (€)	Price per sample (€)
Hardware	CNC 3018 pro	180.00	-
	StarLight, LED ring light, 470 nm wavelength	280.00	-
	Hayear HY-D120X lens	42.00	-
	Olympus EP50 camera	941.00	-
	Metal cabinet	120.00	-
Subtotal		1563.00
Software	(Photoshop annual subscription)	(120.00)	-
	AutoStitch	Free
	Microplastic Annotation Package	Free
	Olympus EPview	Free
Subtotal		0.00 (120.00)
Consumables	AISI 304 filter, ⌀ 47 mm, 10 μm pore size, 100 filter	16.50	0.165
	Nile Red dye, 1 g	295.40	0.0059
	Potassium hydroxide (KOH, purity 85%), 1 kg	8.20	≈ 0.386*
	Hydrogen peroxide (H₂O₂, 30% w/v)	4.60	0.184
	Acetone (purity 99.5%), 1 L	5.60	0.0112
Subtotal		325.70	0.75
Total		2008.70

Consumables, hardware, and software components and their market prices for the detection of microplastics in biota samples using Nile red dye. *The cost per sample for KOH is calculated based on 400 mL of 10% KOH.

Questions about the method?

Nikola Bobchev

Happy to discuss the protocol, the imaging set-up or the data.

✉ bobchev.nikola@gmail.com LinkedIn ↗ ResearchGate ↗