Mould in Geological Collections

Written by Lu Allington-Jones (Principal Conservator), The Natural History Museum, London UK

Mould is probably not at the top of the list of worries of a geological curator, but it does frequently occur in geological collections. It is not only a significant health risk, but it can leave permanent staining, it is an indicator of inappropriately high relative humidity and could spread to other collections.

But how should mouldy geological collections be treated?

Mould

Mould is a micro-organism – a fungus that grows in the form of multicellular filaments called hyphae. It can cause extensive destruction of collagen, lipids and carbohydrates, and hydrolysis of proteinic compounds, leading to embrittlement, loss in tensile strength, staining and deformation of substrates (Valentín 1999, 261). Moulds produce enzymes that split glycerides into glycerine and free fatty acids (Sharphouse 1979,184). They are an extremely diverse and resistant group and under suitable conditions, mould can begin to form within hours. Keeping the relative humidity below 65% will halt the growth of mould (Strang and Dawson 1991), but spores will germinate once favourable conditions are regained. It is difficult to find a treatment that will eradicate spores completely, without causing some damage to the infected substrate. Mould stains may be impossible to remove and spores impossible to eradicate (Guild and MacDonald 2020, 20). The adage that prevention is better than cure becomes particularly resonant.

Above: Mould has grown on this fossil jawbone (NHMUK AQ-PAL-2023-75-PV) in the form of small round white spots. All material coming in from private collections must be checked, especially if they have been stored in attics or garages where conditions may be damp. ©Trustees of the Natural History Museum

Recommended Treatments

Cleaning Geological Specimens

Very fragile specimens can be cleaned with an air puffer and more robust specimens can be dry brushed – both techniques must be undertaken under air extraction or into the nozzle of a vacuum cleaner with a HEPA filter. FFP3 masks must be worn. If specimens are not sensitive to liquids, ethanol can then be applied with a soft brush. A conservator should be consulted regarding wet cleaning techniques, especially when minerals are being treated. Immersion is not advised as this could result in some loss due to pressure changes and small fragments floating away. An excess of solvent could also cause solutes to mobilise and old adhesive to weaken. A pipette can be used to drop controlled amounts of ethanol into cavities or delicate areas that could be damaged by brushing.

Above – Green mould on a hand specimen covering both the matrix and the label. ©Trustees of the Natural History Museum

Above: The presence of mould on the plaster surrounds of this specimen alerted gallery staff to a problem with the fossil marine reptiles mounted on the wall in Victorian frames (specimen NHMUK PVOR40140). Investigations found that the entire wall of specimens was being affected by water caused by a broken gutter, which was seeping from outside through the terracotta. This led to a huge project for conservators and estates staff to remedy the problem and treat the specimens, some of which were deteriorating due to pyrite oxidation. ©Trustees of the Natural History Museum

UV treatment

Although not suitable for treatment of light-sensitive mineral specimens and large items, UV radiation is often a viable option for killing mould on labels which would be sensitive to a liquid treatment. C-type ultraviolet light is the shortest wavelength at 280-100 nm. It is also known as germicidal UV for its use in medical sanitation and the purification of food, water and air. It kills micro-organisms such as viruses, pathogens and mould by destroying the nucleic acids in their DNA. UVC is blocked by Earth’s atmosphere but several UV-generating germicidal devices are available. UVC should be used with extreme care, as exposure to humans can lead to sunburn, skin cancer and blindness. Exposure of objects to UVC should therefore be carried out in a closed environment, with extraction (to disperse the ozone formed), and only accessed after the light has been switched off.

To test the suitability of UVC radiation on labels, small samples can be exposed to the minimum dosage required to kill the mould and then tested for change. Amongst the physical properties of paper, fold endurance is thought to deteriorate the most rapidly, so it is a good indication of the onset of change and is an extremely easy test to do (Padfield 1969). Colour and brightness changes can be assessed by covering part of a label, exposing the rest to UVC, and making judgements by eye. Different mould species require different dosages for annihilation (American Air and Water 2013) so ideally it is important to first identify the mould species present, which can be undertaken by an external company. Alternatively, the most resistant genus likely to be present can be used as a maximum benchmark.

Treating labels

Mould can be cleaned off with a soft brush followed by a smoke sponge or a cosmetic sponge, to remove surface detritus. Each label can then be exposed for the minimum UVC exposure time necessary to kill the most resistant genus present, which is likely to be Aspergillus. The most resistant species in this genus requires 330,000 μWs/cm² for a 99% kill factor (American Air and Water 2013). Treatment time is calculated by converting the wattage of the UV lamp to microwatts and then dividing this by the surface area of the lamp, and then dividing the dosage needed for the worst genus by the answer. e.g. a 55W lamp (usually used for pond water treatment) emits 55 000 000 μW. The surface area of the two 50cm x 1.75 cm bulb cylinders is 1138.04 cm², which means that a dose of 6.83 seconds would be sufficient to kill Aspergillus.

A 55 W compact germicidal lamp can be enclosed in a case and modified by the addition of an on-off switch so dosage can easily be controlled. Prior to treatment of a large number of labels, discolouration can be tested by obscuring half of a label with card before exposure to 7 seconds within the light box. Treatments should only progress if alterations in the optical brightness of the paper and fading of ink are not observed and fold strength is unaffected.

Above: an interesting range of moulds on microscope slide labels. These came from a private collection which had been stored in a garage. ©Trustees of the Natural History Museum

Steps to combat mould

1. Work under extraction, or in a well-ventilated space if this is not available. Cleaning large volumes of mould-contaminated objects indoors without extraction it not recommended. It is essential to wear an FFP3 face mask. A risk assessment should be undertaken and people with respiratory issues or allergies should not attempt carry out this work or be in the same space.
2. Remove all items from the storage box or bag. If this cannot be disposed of this should also be cleaned thoroughly.
3. Lightly brush away the contaminant from each specimen with a soft brush taking care not to damage the surface.
4. If safe for the specimens in question, brush over the surface with 70% ethanol or IMS, as this will kill any living hyphae, and allow to dry naturally.
5. Removing all spores is not possible as they exist everywhere so it is necessary to control the environment and make growth unlikely. Monitor the environment, keep the RH lower than 65%.
6. If a plastic box is being used for re-storage, a buffer material, such as prozorb silica gel, must be placed inside, to help stabilise the relative humidity. This will help prevent further mould growth.
7. Also useful are RH indicator cards

• Confine it
• Stop growth
• Eradicate it
• Prevent it from happening
• Prevent it from entering the collections

Rationale behind rejection of other treatments

Freezing

CCAHA (2013) recommend freezing bagged specimens before drying them out, as a first step in disaster management following an outbreak of mould within collections of historic artefacts. Drying must then be undertaken because the water is liberated upon defrosting, and available to react with the mould. If freezing is used as a temporary quarantine measure, it is advisable to wrap specimens in polythene, incorporate absorbant materials and allow the enclosures to acclimatise to ambient temperatures for 24 hours before opening. This will reduce the potential for rapid changes in relative humidity and also condensation, which could cause the mould to flourish. Freezing (and low-oxygen environments) will limit mould growth by sending it into a dormant state and it will kill hyphae, but it will not kill the spores. Some species, e.g. Aspergillus niger, will even flourish down to -10^oC, whilst other species are anaeorobic (Valentín 1999, 261). Freezing is also not appropriate for some geological material so it is better to clean the mould off immediately rather than undertaking freezing.

Drying

If you dry out mouldy specimens, the mould can re-activate if humidity rises again. Freeze-drying has, however, been used in paper conservation to halt deterioration. The freeze-drying process involves temperature reduction of objects so that all water is converted to ice and then the ice is evaporated by sublimation in a vacuum. Parker (1993) used freeze-drying on vellum and found that dimensional changes were significant during the process but decreed that this was acceptable for the subject material. Some geological material will not be adversely affected by freeze-drying, but others with suffer permanent damage (such as minerals with fluid inclusions as well as material containing soluble salts or clays). It is best in general not to freeze-dry geological material.

Gamma Irradiation

Ionising radiation does not leave any chemical residue behind, and will not affect inks, so it initially appears to be a suitable treatment for labels. Studies have found that, at certain doses it is 100% effective at killing mould on paper, whilst dry and wet cleaning methods and ozonation often only reach 50% eradication (Sinco 2000, 40). Whilst some reports state damage was ‘minimal’ at 12 kGy, others found up to 100% accelerated aging (embrittlement and yellowing) of paper at 10 kGy (Sinco 2000, 38; 39). Adamo et al (2001) found that at 5 kGy gamma irradiation significantly reduced the whiteness of paper by both increasing the light absorption coefficient and reducing the ISO brightness. It also caused depolymerisation of cellulose, the formation of unsaturated compounds and yellowing (the latter manifesting after artificial aging experiments). Irradiation causes the formation of free-radicals in cellulose which will react with oxygen and decreases polymerisation. At 2 kGy damage to paper was insignificant (Adamo et al 2001) but doses lower than 10 kGy are ineffective for eradicating mould (Sinco 2000, 39). Radiation is seen as a disaster mitigation technique but it is recognised to cause further damage. Radiation can also cause colour changes in some minerals and is not recommended in general.

Other types of wet cleaning

Historical attempts to kill mould include: sodium ortho-phenyl-phenate, hydrogen cyanide, sodium pentachlorophenate, isopropyl metacresol, formaldehyde, DDT, methyl bromide, p-chloro-m-cresol, ethylene dioxide/oxide and ammonium salts (Woods, 2006). Wagstaffe and Fidler (1968) recommend mist-spraying bird taxidermy with a 10% alcoholic solution of thymol and treating the legs, feet and bill by painting them with Mystox, phenol or formalin. Most of the treatments mentioned could cause discolouration, drying and pose health risks (Voronina et al 1980; Woods 2006). Commercially available fungicides should be avoided because their long-term effects are unknown and they may leave residues on the specimen. South East Museums (2019) recommend cleaning with surgical spirit, but state that a conservator should be consulted first, to check that the material is suitable. 

Voronina et al. (1980) tested a quaternary ammonia compound on parchment: alkyldimethylbenzylammonium chloride (Catamin AB) at 3% in 1:3 water-alcohol solutions. This worked as a fungicide and bactericide when parchment was immersed for 5 minutes, the solution was brushed on, or if the object was placed between impregnated filter paper for 40 minutes. This technique would be unsuitable for geological specimens, but could be applicable to the labels, if the inks are not affected by any of the components and if the paper is not acidic. Cationic detergents such as this, although possessing antiseptic properties, are strongly attracted to acidic compounds because they are negatively charged. This makes residues difficult to remove after treatment (Rivers and Umney 2003, 535).

Guild and MacDonald (2020, 22; 29) recommend only dry brushing but make no suggestions for killing ingrained spores, which remain a health risk and can reactivate under suitable conditions. CCAHA (2013) suggest the use of vulcanised rubber sponges after vacuuming, but this is only suitable for items with a smooth surface. Mason (1989) suggests vacuuming, brushing and a “light” application of isopropyl alcohol for leather, but only dry cleaning is recommended for textiles (CCI 2016). Strang and Dawson (1991) recommend the use of ethanol as a fungicide for skins and furs. This is suitable for most fossils and rocks, but care should be taken with some minerals.

Further Reading:

Didem Aktas, Y., Shi, J., Blades, N. and D’Ayala, D. 2018. Indoor mould testing in a historic building: Blickling Hall. Heritage Science 6: 51.

Florian, M-L. E. 2002. Fungal facts: solving problems on heritage collections in museums and archives. London: Archetype.

Pack, C. A. 2011. A fungus among us: mold growth in museum environments. https://citeseerx.ist.psu.edu/document?repid=rep1&type=pdf&doi=cb32d62d3b49441d1f8fcca535598401e3f2eff1

Thacker, C. E., Feeney, R., Camacho, N. A. and Seigel, J. A. 2008. Mold removal and rehousing of the ichthyology and herpetology skeletal collections at the Natural History Museum of Los Angeles County. Copeia (4): 737-741.

References

Adamo, M., Brizzi, M., Magaudda, G. 2001. Gamma radiation treatment of paper in different environmental conditions: chemical, physical and microbiological analysis. Restaurator, 22(2), 107-131.

American Air and Water. 2013. UV irradiation dosage table. http://www.americanairandwater.com/uv-facts/uv-dosage.htm Accessed 17 December 2013.

CCAHA. 2013. Managing a mold invasion: guidelines for disaster response. Conservation Center for Art and Historic Artifacts. http://www.ccaha.org/uploads/media_items/managing-a-mold-invasion-guidelines-for-disaster-response.original.pdf. Accessed 3 January 2014.

CCI. 2016. Mould Growth on Textiles. CCI Notes 13/15. Ottawa: Canadian Conservation Institute.

Guild, S. and MacDonald, M. 2020. Mould Prevention and Collection Recovery: Guidelines for heritage collections. Ottawa: Canadian Conservation Institute. https://www.canada.ca/en/conservation-institute/services/conservation-preservation-publications/technical-bulletins/mould-prevention-collection-recovery.html

Mason, J. 1989. Removing mould from leather. CCI Notes 8/1. Ottawa: Canadian Conservation Institute.

Padfield, T. 1969. The deterioration of cellulose, In: Problems of Conservation in Museums, International Council of Museums, Publication number 8, Eyrolles, Paris, 119-164.

Parker, A. E. 1993. Freeze drying vellum archival materials. Journal of the Society of Archivists. 14 (2), 175-188.

Rivers, N. and Umney, S. 2003. Conservation of Furniture. Butterworth-Heinemann, Oxford.

Sharphouse, J. H. 1979. Leather Technician’s Handbook. Leather Producer’s Association: London.

Sinco, P. 2000. The use of gamma rays in book conservation. Nuclear News 43(5): 38-40.

South East Museums. 2019. Mould. http://southeastmuseums.org/mould#.WaxAOBmGPrc Accessed 3 September 2017.

Strang, T. J. K. and Dawson, J. E. 1991. Controlling museum fungal problems. Canadian Conservation Institute Technical Bulletin 12. Ottawa: Canadian Conservation Institute.

Valentín, N. 1999. Evaluation of microbial contamination in proteinaceous materials: control methods. In: R. Larsen (ed.) Methods in the analysis of the deterioration of collagen based historical materials in relation to conservation and storage. Preprints of the Advanced study course 1999, 6-10 July. Copenhagen : School of Conservation, Royal Danish Academy of Fine Arts. 261-265.

Voronina, L., Nazarova, O. and Petushkova, Y. 1980. Disinfection and straightening of parchment damaged by micro-organisms. Restaurator 4(2), 91-98.

Wagstaffe, R. and Fidler, J. H. 1968. The preservation of natural history specimens: Volume II. H F & G Witherby Ltd, London.

Woods, C. 2006. The conservation of parchment. In: M. Kite & Thomson, R. (eds) Conservation of Leather and Related Materials. Elsevier, London. 200-224.