Jellyfish envenomations, which have been increasing worldwide, result in myriad clinical outcomes, ranging from sting-site pain and inflammation to life-threatening sequelae and death [1
]. In response to heightened public health concerns and lay inquiries, clinicians and emergency-care personnel often rely on tertiary Internet resources, such as Medscape, eMedicine Health, or Mayo Clinic Online [2
]. These clinical management summaries cite an assortment of primary peer-reviewed references, many of which are contradictory and/or uncorroborated [5
]. Further confusion results from authoritative Internet articles advising both clinicians and the general public [8
], which often echo and extrapolate from selected studies without critically discussing pertinent limitations or divergent results. For example, recent articles have uncritically perpetuated already-hyperbolic press releases based on small-scale, in vitro
model studies [9
] into headlines declaring how vinegar “can be deadly” and “may kill” [10
], suggested disproven treatments like ice packs [8
], and recommended the use of shaving cream [8
] and/or sting-site scraping, which have never been corroborated by any human study or validated model [13
] and involve the application of site pressure which has been shown to worsen outcomes. The confused lay press recommendations and lack of validated protocols are especially dangerous in the case of certain cubozoan envenomations, which cause more loss of life than shark attacks annually [15
Rigorous, non-redundant and mechanistically distinct testing of the efficacy, as well as the potential harmful side effects, of many jellyfish treatment options is important because animal models indicate that the pathology is venom dose-dependent and, thus, if treatment methods increase venom dose or “load”, they may cause more harm than good [19
]. Additionally, since emergency responders may not have standardized approaches available, especially in remote areas, it is important to know what other options cause the least harm.
However, because jellyfish envenomations, and particularly cubozoan envenomations, are potentially dangerous, live tentacle application involving human participants to comparatively assess different first-aid measures is inherently hazardous. While the potency of the tiny carybdeid, Carukia barnesi
, to cause Irukandji syndrome, used investigator self-testing and self-consenting tests [22
], broader human studies are ethically problematic. Mouse and piglet animal models have been extremely valuable in the elucidation of pathogenic mechanisms [19
], but low cost and reproducible ex vivo
and in vitro
laboratory-based models are critically needed to accelerate progress and rigorously investigate approaches to mitigate cnidae discharge and venom activity in the determination of best practices. Ideal models would be broadly applicable, reproducible, easy-to-measure, and accurately recapitulate sting events. To defensibly correlate to the biological reality of an authentic sting, ideal models would meet the following criteria:
(1) Exhibit spontaneous tentacle cnidae discharge. We observed that application of live, freshly-cut tentacle sections onto low melting point agarose gel sections made up with fresh, intact human red blood cells (RBC) elicited an immediate potent sting response with discharged nematocyst tubules visibly penetrating deep into the blood agarose, as well as tentacle contraction, indistinguishable from spontaneous stinging after application of tentacle sections on human skin [23
]. Artificial stimuli can induce explosive sting-type events in which essentially all venom-filled penetrant cnidae (nematocysts) discharge or non-productively rupture. Specifically, electrical (DC voltage) stimulation causes massive circumferential tentacle nematocyst discharge, as well as visible tentacle damage with heat effects, and is in extreme excess of physiological electrical excitation [24
]. Chemical stimuli, such as exposure to alcohols, can also cause nearly all tentacle nematocysts to explosively fire venom-delivering hypodermic-like nematocyst tubules [26
]. But in both cases, nearly all cnidae are fired and other tentacular cell types are disrupted or lysed. Thus, the toxic exudate from either approach differs markedly from authentic sting-associated venom both quantitatively and qualitatively.
(2) Allow for visualization and quantification of cnidae discharge. Envenomations involve the firing of up to thousands of packed cnidae per linear tentacle unit of length. It is critical that potential first-aid approaches to remove adherent tentacles be tested to examine whether a specific approach reduces or induces additional cnidae discharge. An optimal envenomation model must allow for imaging and ideally quantification of cnidae discharge with which to experimentally assess test conditions (e.g., topical temperature or pressure) or test solution efficacy to prevent, inhibit, or exacerbate tentacle cnidae discharge.
(3) Measure venom activity directly. An ideal model of envenomation would allow direct and time-dependent determination of venom activity in the experimental tissue model. The hemolytic activities of jellyfish venoms have been well-studied, and hemolytic units (specific activity per venom mass) for different species have been calculated [20
]. Such biochemical baselines provide clear standards for envenomation models. Experimental models that do not result in levels of venom activity comparable to primary or laboratory purified venom samples are not credible proxies for sting events.
(4) Employ rigorous controls. It is also critical to investigate the effects of comparative controls including null treatment or the application of a control “mock” solution such as seawater to exclude experimental artifacts, including direct physical manipulation effects of adding solutions in treatment activities. Such controls are especially important given that there is no careful study to determine what percentage of cnidae discharge during natural sting events; far less than 100% of the cnidae present on a tentacle may actually fire. It is also critical to assess converse interpretations of results, such as whether a tested solution may enhance venom recovery from tissue and thus reduces “venom load”, or alternatively, whether site pressure alone from the application of a treatment causes or enhances venom expulsion from already-discharged nematocysts [27
]. In the later example, the amount of venom injected simply by the application of the first-aid measures or sting-site “scraping” should be compared to the venom injected and potentially injected without such measures in efforts to ascertain treatment choice and effectiveness.
Previous studies have sought to test first-aid measures in a diversity of cnidarian species [28
]. However, these experiments varied widely in methods and results, and none employed comprehensive approaches, which met the four above-mentioned criteria simultaneously. In this study, we have employed an array of experiments to rigorously test first-aid approaches from multiple angles. In addition, we unveil a novel ex vivo
model, which alone is able to meet the four requirements listed above. Apart from satisfying these criteria, the model is simple and inexpensive and, thus, could be employed in laboratories with minimal budgets and limited equipment.
3. Experimental Section
In an effort to develop methodologies that can be employed broadly in other laboratories with other cnidarian species, a tiered research approach was designed to examine a broad array of commonly used first-aid measures, as well as commercial preparations (see Table 1
and Table 2
). Instead of forcing nematocysts to fire, our novel models capitalize on the spontaneous stinging response of live tentacles in the presence of blood cells. Venom activity was measured by hemolysis of live blood cells, thus using a well-documented primary venom activity, while more accurately recreating the natural stinging action that occurs during an envenomation event.
The solutions used in all assays are as follows: seawater (Instant Ocean formulated at 35 ppt; Spectrum Brands Inc., Backsburg, VA, USA), freshwater (double-distilled), vinegar (white distilled; Bakers and Chefs CJ314, SAM’s West Inc., Bentonville, AR, USA), lidocaine (4% in 150 mM saline; MP Biomedicals LLC, Solon, OH, USA ), ethanol (Pharmco-Aaper, Brookfield, CT, USA), isopropanol (Fisher Scientific, Fair Lawn, NJ, USA), Epsom salts (CVS Pharmacy Inc., Woonsocket, RI, USA, used saturated in double-distilled water), copper gluconate (30 mM in 150 mM saline; Strem Chemicals, Newburyport, MA, USA).
3.1. Tentacle Solution Assay (TSA) to Evaluate Nematocyst Discharge
To determine the effects of first-aid treatments on nematocyst discharge, 15 μL of each solution was added to a coverslip-bottomed slide well containing a 5 mm piece of live, freshly cut tentacle (for a complete list of treatments tested, see Table 1
). Video and still images were recorded using a dissecting microscope. For pre-treatment experiments, tentacles were allowed to incubate in the solution for 30 min. In a separate set of wells, tentacle pieces were pre-treated with 15 μL of test solutions for 1 min. The test solution was then removed and 15 μL of isopropanol was added to determine if the solutions irreversibly prevented cnidae discharge.
3.2. Ex Vivo Assays to Evaluate Hemolytic Activity
We utilized live human RBC from normal donors (approved protocol CHS#12561, University of Hawaii Committee on Human Studies) and low melting point agarose to constitute a live red blood cell agarose to measure hemolysis, a well-documented venom activity [19
]. To create the blood agarose, fresh human RBC were washed and resuspended in modified RPMI (“YRPMI”: 23.81 mM NaHCO3
, Fisher; 102.67 mM NaCl, BDH; 5.37 mM KCl, Fisher; 0.41 mM MgSO4
O, Fisher; 25 mM HEPES, Fisher; 6.67 mM NaH2
, Fisher; 0.42 mM Ca(NO3
O, Fisher) at 3% and kept at 37 °C. Low melting point, molecular grade agarose (Nusieve GTG Agarose, Lonza, Rockland, ME, USA) was dissolved in YRPMI at 60 °C then cooled to 39 °C. Equal volumes were mixed (final concentrations: 1.5% RBC, 1.5% agarose in YRPMI) and the RBC-agarose solution was immediately aliquotted onto glass to form uniform rectangles as desired and placed at 30 °C to gel. Slides were then maintained in a humidified tissue culture incubator at 37 °C within 4% CO2
The same blood agarose was used for the Venom Blood Agarose Assay (VBAA) and Tentacle Blood Agarose Assay (TBAA). Since some test solutions exhibited direct pH and osmotic effects, we also designed a “skin” layered blood agarose model (Tentacle + Skin Model Blood Agarose Assay or “TSBAA” model). This model also initiated spontaneous stinging in live tentacles. Live time microscopic examination of both tentacle model systems revealed that nematocyst tubules forcibly ejected into the blood agarose layer to result in localized clear nascent zones of lysis of live human RBC. The objective was to develop a reproducible model to recapitulate authentic envenomation amendable to direct assessment of “venom load” or dose via quantification of a known venom activity: hemolysis.
3.2.1. Venom Blood Agarose Assay (VBAA)
Wells 2 mm in diameter were made in 2 mm thick agarose using a biopsy punch (Sklar Instruments, West Chester, PA, USA). Five microliters of venom or diluted venom (serial dilution using 0.5 M citrate) was added to each well and incubated at 37 °C. Photos of lytic zones were taken at 1 h and overnight. The volume of the lytic zone was calculated using the three radii in ImageJ (U.S. National Institutes of Health, Bethesda, MD, USA) used to determine the volume of the cylindrical ring of lysis around the well (cylinder volume minus well volume). These data were then used to determine dose response curve seen in Figure 2
using known hemolytic units [19
], the linear portion of which was determined for the calculations in Figure 3
3.2.2. Tentacle Blood Agarose Assay (TBAA)
Approximately 1 cm of freshly cut tentacle was applied to each slide: tentacles were weighed after the experiment. Slides were then incubated at 37 °C. Images were recorded using a high-resolution scanner (Epson Perfection V500 Photo Scanner, Long Beach, CA, USA), dissecting- or inverted- microscope (Olympus model SZX16 or CKX41SF, Olympus Corporation, Tokyo, Japan) at specific time points. The area of the zone of hemolysis was calculated using ImageJ. Briefly, scale was set using the known slide width and 15 mm × 15 mm subsections were taken from each slide for analysis to remove edge effects. Control slides for each solution (solutions applied without tentacles) were used to set the color threshold for no hemolysis. The percent area of the hemolytic zone was taken directly from the “analyze particles” function. Zone of hemolysis was calculated as the percent total area lysed (Figure 4
3.2.3. Tentacle Skin Blood Agarose Assay (TSBAA)
Because vinegar and other potential agents led to direct osmotic- or pH-based hemolysis and to better recapitulate the tissue layers involved in authentic tentacle envenomation, we designed a modified blood agarose model with a protective skin layer. Blood agarose slides were prepared as described in the TBAA model but overlaid with a “skin” comprised of pig small intestine modeled after previous published methods for preparation of skin grafts [56
]. Specifically, porcine small intestine sections were rinsed in 50 mM saline and sectioned and cut along their length to create thin, flat sheets. Sections were sterilized in 5% Hydrogen peroxide and 10% ethanol in 50 mM saline for two hours, then rinsed in sterile YRPMI three times for 30 min each. Intestinal sections were then stretched over glass to create a thin, flat sheet and allowed to air-dry. The resultant membrane was rubbed with pharmaceutical grade anhydrous lanolin and placed atop the blood agarose squares; these could be removed easily.
In the pre-treatment condition, creams were applied to both sides of the skin and allowed to penetrate for ten minutes. Pre-treated skins were then applied to agarose squares, and approximately 1 cm of tentacle was allowed to spontaneously sting for five minutes.
To test efficacy as tentacle removal solutions, fresh tentacles were allowed to sting for three minutes before 50 μL of the test solution was applied directly onto the tentacle. The treated tentacles remained for another two minutes (for a five-minute total sting time) before the skin was removed.
Lastly, to test efficacy as post-sting treatments, tentacles were allowed to sting spontaneously for five minutes, and 50 μL of sting treatment solutions were added to the skin after tentacle removal. The post-treated skins were removed after two minutes.
All slides were placed at 37 °C for one hour, and the area of the zone of hemolysis was calculated using ImageJ as described for the TBAA (Figure 6
). Significant differences between treatments were tested using one-way ANOVAs with Holm-Sidak planned multiple comparison tests to examine differences between tested treatments and no treatment [57
The recent debate about the efficacy of vinegar, based on the study by Welfare and co-workers [45
], and the related press release-extrapolated claim that vinegar application increases venom load and thus “can kill” demonstrates how poor experimental design models which lack appropriate rigorous controls, would fail to meet fundamental validation criteria. By using newly developed ex vivo
envenomation models, we have demonstrated that vinegar inhibits nematocyst discharge and that copper gluconate-based products (Sting No More™) inhibit hemolytic activity subsequent to stings of the cubozoan Alatina alata
when used before, during, or after a sting event. Common lay approaches, including urine, were not significantly better than seawater. Based on these data, we conclude that current first-aid protocols for jellyfish stings, including the removal of tentacles by the application of vinegar, do prohibit further cnidae discharge and do not elicit adverse outcomes. Lidocaine has been shown in previous studies to slightly lessen pain and to inhibit nematocyst discharge [33
]; that was not confirmed in this study. These data also suggest that while vinegar can and should be used as tentacle-removal solution as it prevents both chemically- and pressure-induced cnidae discharge, it should not be considered an effective treatment for the sting itself. Because of the profound cnidae discharge induced by ethanol and isopropanol and their failure to reduce hemolysis when applied to adherent tentacles, these data do not support their use as treatments. The potent inhibitory effects of copper gluconate-based products (Sting No More™) to hemolytic activity subsequent to stings of the cubozoan Alatina alata
stings when used before, during or after a sting event provides a promising new therapeutic tool for the treatment of cnidarian envenomation.