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Review

Lipid Analysis by Thin-Layer Chromatography—Detection, Staining and Derivatization

by
Johanna W. Schubarth
†,
Jenny Leopold
†,
Kathrin M. Engel
† and
Jürgen Schiller
*
Institute for Medical Physics and Biophysics, Faculty of Medicine, Leipzig University, Härtelstr. 16-18, 04107 Leipzig, Germany
*
Author to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Lipidology 2026, 3(1), 3; https://doi.org/10.3390/lipidology3010003
Submission received: 9 October 2025 / Revised: 5 November 2025 / Accepted: 5 January 2026 / Published: 13 January 2026
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Abstract

Thin-layer chromatography (TLC) remains a widely used, cost-effective and convenient method to separate small molecules, particularly in the field of natural products and (phospho)lipids. Despite advances in chromatographic methods such as high-performance liquid chromatography (HPLC), TLC retains several advantages, including simplicity and accessibility. However, a critical step is the visualization of the separated lipids on the TLC plate. Although the majority of the regularly used methods were established decades ago, there are still a number of potential pitfalls and widely unknown aspects. This review provides a concise overview about commonly used stationary phases and the solvent systems in TLC analysis of lipids. The main focus is on visualization techniques, spanning from non-specific, destructive (charring by semi-concentrated acids) to specific, non-destructive approaches (e.g., exposition to iodine to monitor unsaturated lipids). The advantages and disadvantages of the different methods will be critically discussed and frequently occurring problems highlighted. Furthermore, the combination of TLC with mass spectrometry (MS) detection will be introduced, covering both extraction-based electrospray ionization MS techniques as well as desorption techniques such as matrix-assisted laser desorption/ionization MS. MS detection, while generally more sensitive and offering molecular specificity, introduces higher technical and financial requirements compared to conventional staining. Nonetheless, the combination of TLC with MS holds significant potential for enhancing lipidomic workflows, particularly in complex biological samples.

Graphical Abstract

1. Introduction

Although high-performance liquid chromatography (HPLC), and particularly gas chromatography (GC), offer higher separation quality, classical thin-layer chromatography (TLC) and its advanced version, high-performance thin-layer chromatography (HPTLC), are also widely used, particularly in the field of natural products and lipid analysis. The most important differences between TLC and HPTLC are (a) the different particle sizes of the stationary phases (about 10–12 µm in TLC and 5–6 µm in HPTLC), (b) the thickness of the stationary phase (about 200 µm in TLC and 100 µm in HPTLC) and (c) the care that should be taken when applying the samples and processing the obtained data [1]. The most important advantages of planar chromatography in comparison to HPLC can be summarized as follows [2]:
  • Since several samples can be analyzed in parallel, i.e., simultaneously, TLC is typically faster than HPLC.
  • TLC is a simple method, and the equipment is relatively inexpensive. It can be easily installed and established in virtually all laboratories. HPTLC, however, relies much more on automated procedures and is, thus, more expensive.
  • Only minimal maintenance is required, as there is hardly any risk that devices will need repairs. HPLC columns, in contrast, become easily plugged and/or contaminated.
  • TLC requires only small solvent volumes. Therefore, it is considered more sustainable compared to HPLC, which consumes much larger volumes of solvents.
  • There are absolutely no memory effects, since a new stationary phase (i.e., a new TLC plate) is always used. In contrast, the HPLC column is used many times and flushed intensively to remove residual components from a previous run, yet contamination may remain in the column.
  • Chromatograms can be visually evaluated directly on the plate, and “digital images” of the plates can be obtained, processed, and saved. These images are often more illustrative than “pure” HPLC data.
Notably, the significant success of TLC with many applications only became possible with the commercial availability of industrially manufactured TLC plates with significant homogeneity: the (in the majority of cases) silica gel layer on earlier home-made TLC plates was never very homogeneous and, thus, the achievable separation quality was comparably poor.
Although reversed phase (RP) columns are commonly used in HPLC, normal phase (NP) is most commonly used for TLC separations. Accordingly, non-modified silica gel is the stationary phase of choice in TLC and is also used in the majority of lipid separation problems. This is mainly due to historical reasons and the fact that RP HPLC is compatible with aqueous samples and biomolecules such as proteins [3]. Since lipids are apolar, NP is typically used as the standard approach. Although not the focus of this review, an additional feature of TLC has to be mentioned: preparative TLC, which may be performed, e.g., to isolate small quantities of lipids (up to a few milligrams) [4]: the silica gel representing the spots can be scraped off and the lipid eluted for use in further experiments, such as the detailed structural analysis of novel or unknown lipid classes. Dedicated TLC plates with enhanced silica layer thicknesses are commercially available for this purpose [5].
We will discuss the different steps of TLC only to a very limited extent (practical tips are available in [6]). The most important step of lipid analysis by TLC is the extraction (and, thus, enrichment) of lipids from the body fluid (e.g., blood) or the tissue of interest. Although there is growing opposition to the use of chloroform due to its toxicity, extractions with chloroform and methanol (originally published by Folch and coworkers [7] and with slight modifications by Bligh and Dyer [8]) are still the most widely used. The reason for this is that it typically produces the highest lipid yields—although this strongly depends on the system of interest, particularly its water content. However, this is only valid for the bulk lipids such as phosphatidylcholine (PC) and phosphatidylethanolamine (PE), i.e., the most abundant phospholipids in the majority of biological samples. The complete extraction of more polar lipids such as glycolipids or phosphoinositides (with several phosphate groups and, thus, charges) is considerably more challenging. In these cases, increased ionic strength and/or a decreased pH is necessary for the screening of the charges in order to improve the extraction yield. The topic of lipid extraction was recently comprehensively reviewed [9].
There are many established solvent systems for the successful separation of (phospho) lipids. We (and many others) typically use a solvent system of chloroform, ethanol, water and triethylamine (30:35:7:35, v/v/v/v) because it works best, in our opinion, for phospholipids [10]. Apolar lipids such as cholesterol and triacylglycerols require less polar solvents [11].
The migration of a lipid in/on the TLC plate is basically determined by two different parameters:
i.
The interaction with the silica gel (normally unmodified silica gel) on the TLC plate;
ii.
The interaction of the lipid with the solvent mixture.
The more marked the interaction with the stationary phase, the smaller the migration distance of the lipid and vice versa. Thus, small Rf (“ratio of fronts”) ratios indicate a strong interaction of the analyte with the silica gel.

2. Making the Analyte Visible

Basic chemistry textbooks often describe the separation of selected mixtures using examples such as leaf pigments [12] or inks. This is not normally due to the huge interest which these compounds are attracting, but rather the fact that these compounds are visible already without any pretreatment, as they are colored and absorb visible light (λ ≈ 380–780 nm). This approach is, of course, not applicable to “native” lipids because they do not contain chromophores which would render them visible without suitable staining. The structural fragments that are typically targeted during staining, such as the double bonds within the acyl residues, the phosphate and the headgroups, such as choline or ammonia, are illustrated in Figure 1.
Since all naturally occurring lipids are colorless, the developed TLC plates have to be stained in order to render the different lipid classes visible. A lot of different ready-made reagents are commercially available nowadays (for instance, from MERCK), and there is often no need to prepare the reagents in the laboratory. Comprehensive reviews of commonly used dyes are available in [13,14].
Since the majority of organic stains are relatively apolar (i.e., insoluble in water), they tend to become enriched in the apolar phase, i.e., in the lipid spots on the TLC plate. Consequently, the binding between the dye and the lipid is, in many cases, non-selective. The addition of weak acids such as acetic acid or salts (e.g., NaCl) additionally increases the polarity of the aqueous phase and reduces the solubility of the dye. This shifts the dye to the lipid phase and intensifies the staining of the lipid spots. Different non-specific dyes (such as Congo Red, Bromthymol Blue, Ponceau S and many others) were recently compared regarding the achievable sensitivity towards the staining of lipids [15].
The discussion of the different approaches, their advantages and drawbacks as well as potential pitfalls will be discussed in the next section. All relevant dyes can be sorted according to their specificity and whether they are destructive or non-destructive.

2.1. Destructive, Specific Dyes

Dyes listed here are characterized by a covalent modification of the analyte. This requires the reaction with a particular functional group, such as an amino group. However, it must be noted that this definition is not universally applied, and some authors sort staining protocols differently.
It is textbook knowledge that there are some chemicals which react with defined functional groups of the analyte molecules under generation of colored products, often accompanied by a change in the molecular weight. The best-known example is presumably the derivatization of carbonyl groups (such as aldehydes) by 2,4-dinitrophenylhydrazine, which is also useful for the detection of oxidized lipids [16]. Another suitable agent for the detection of truncated oxidized aldehyde lipids is 1-pyrenebutyric hydrazide (PBH) [17], which was recently applied and gives excellent sensitivity. Lipid oxidation is often accompanied by the scission of a double bond and the formation of the corresponding aldehyde as the primary oxidation product. An excess of the oxidant leads to the formation of the respective carboxylic acid as a secondary oxidation product [16]. Since “standard” lipids do not contain any reactive carbonyl groups, other staining strategies are necessary in order to detect non-oxidized lipids.
A survey of frequently used staining agents is available from the comprehensive internet website www.cyberlipid.org (accessed on 5 November 2025), which is dedicated to all aspects of lipid chemistry. Additional overviews can be found in the excellent review by Sherma and Bennett from 1983 [18] and a more recent version [19]. Another review with a focus on the investigation of petroleum products is available in [20].
Some selected staining approaches are summarized in Table 1. Only a few representative methods will be discussed in more detail in addition to the dyes listed in Table 1.
One very sensitive staining reagent is 0.2% amido black 10B (“Naphthol Blue Black”) in 1 M NaCl: after pre-soaking in water, the TLC plate is immersed in the staining solution for a few minutes. The sensitivity achieved is approximately 15 ng regarding DAG, TAG and PS, which are the most sensitively detectable lipid classes using this staining protocol. In contrast, only about 100 ng of free fatty acids and 500 ng of phorbol esters can be detected. There is an ongoing debate about whether dipping and spraying methods truly yield identical results [21].
Ceric ammonium molybdate staining was recently used to visualize lipids on the TLC plate and detects lipids in amounts of about 130 ng. However, this method provides only semiquantitative data, since staining intensity is significantly affected by the degree of fatty acyl unsaturation [22]. Ninhydrine staining can be used for lipids but is limited to lipids with primary amine functionality, such as PE and phosphatidylserine (PS).
Table 1. Selected widely used staining protocols for lipids.
Table 1. Selected widely used staining protocols for lipids.
MethodCharacteristics/SpecificityReference
NinhydrineFor the detection of lipids with amino groups (phosphatidylserine (PS) and -ethanolamine (PE)) as reddish spots. Ninhydrine staining may be combined with primuline staining to detect all PL classes.[23]
1,6-diphenyl-1,3,5-hexatrieneA two-step staining protocol. Initially, developed TLC plates are sprayed with the lipophilic fluorochrome 1,6-diphenyl-1,3,5-hexatriene, followed by application of the phosphorus-specific molybdenum blue (Dittmer–Lester) reagent (vide infra). Detection limits of down to 10 ng phospholipid are achieved, which is a considerable sensitivity enhancement compared to the individual dyes.[24]
Dragendorff’s reagentA complex mixture containing basic bismuth nitrate that exclusively stains choline-containing lipids, particularly PC, lysophosphatidylcholine (LPC) and sphingomyelin (SM).[25]
Molybdenum blue reaction according to Dittmer and LesterDetects phosphate in phospholipids through a chemical reaction involving molybdenum blue chemistry. The reagent contains ammonium molybdate, which reacts specifically with phosphate ester groups, which reduce molybdate ions to form a colored complex known as molybdenum blue.[26]
Ceric ammonium molybdate (CAM)Mo (VI) oxidizes unsaturated lipids into carbonyl compounds on the HPTLC plate upon heating, while itself being reduced to Mo (IV).[22]

2.2. Non-Destructive, Non-Specific Dyes

A comparably simple and established staining method for lipids is the exposure of the developed TLC plate to elementary iodine (I2). Solid I2 is always accompanied by violet-colored vapor. This means that I2 undergoes sublimation, i.e., a transition at ambient conditions (standard temperature and pressure) directly from the solid phase to the gas state. Gaseous I2 reacts directly with the lipids on the TLC plate under generation of a non-covalent, brown complex representing the additional product of iodine. I2 is presumably only loosely bound [27] and the intensity of the stained spot decreases in a time-dependent manner because I2 evaporates. Thus, for further analysis of the analyte, I2 can be removed, exposing the TLC plate to a slight vacuum. This removal is not quantitative, since there is an equilibrium between “bound” and “free” I2.
Another problem regarding quantitative analysis is the dependence of the staining intensity on the double bond content of the samples: saturated (phospho)lipids can only be marginally stained, whereas highly unsaturated lipids (particularly with docosahexaenoyl residues, i.e., with six double bonds) are overestimated. In general, the more double bonds, the greater the I2 binding and the staining intensity on the TLC plate. Lipid classes like PC and PS can only be scarcely detected under UV after exposure to I2 vapor [28], although this behavior is not yet well understood. In contrast, PE binds I2 the most efficiently [27]. The exact molecular basis of I2 binding to unsaturated lipids remains unclear, and a mechanism such as lipid “entrapment” like starch–I2 complexes could be proposed.
TLC plates impregnated with silver nitrate (AgNO3) are often used to separate lipids with varying degrees of unsaturation. With increasing double bond content, more silver ions are bound, slowing down the migration of the respective lipid on the TLC plate. It was shown that polyunsaturated lipids exhibit intense darkening when the separation was performed on AgNO3-impregnated TLC plates [29]. This can be explained by the reduction of Ag+ to colloidal silver. Remarkably, this darkening is dependent on the composition of the solvent system and seems to require the presence of aromatic hydrocarbons such as toluene [30].
Coomassie brilliant blue staining is also widely known from protein analysis (e.g., the “Bradford” assay) but can also be applied as a method to detect lipids on TLC plates [31]; the visual detection limits of this stain for lipids (cholesterol, cholesterol esters, glycerides, phospholipids, ceramides and glycosphingolipids) range from about 0.05 to 0.5 µg. Recently, Coomassie brilliant blue staining was combined with MS detection [32], which emphasizes that the molecular weight of the lipids remains constant upon the reaction with Coomassie brilliant blue.
Some selected staining approaches are summarized in Table 2.
Staining with 2,7-dichlorofluorescein or rhodamine 6G [34] provides yellow or pink spots, respectively, if the TLC plate is observed under UV light. Both dyes can be removed from the lipid spots if the polarity of the solvent is changed or the lipid (with the bound dye) is passed over a short column. This also applies for the dye primuline [42], which can be used in a similar way and yields sensitivities in the low nanomole range [23]. This sensitivity is comparable to staining with rhodamine [34].

2.3. Destructive, Non-Specific Dyes

Spraying the entire TLC plate with a corrosive reagent, such as sulfuric acid, and charring the plate (150–200 °C) to render the lipids visible as black spots is an established, widely used method [43] but nowadays considered somewhat crude [44].
A commonly used approach employs 50% sulfuric acid (H2SO4) either in methanol or water, after which the developed plate is heated to approximately 120 °C for about 60 min. The charring is caused by the loss of water due to the strong dehydrating agent H2SO4. The generation of elementary carbon from lipids (as in the case of carbohydrates if treated with sulfuric acid) was also used to explain the formation of colored spots [45]. Although some details are still unknown, it should be noted that saturated and unsaturated lipids require different times for complete reduction to carbon. The intensities of the obtained black spots are useful data to determine the absolute amounts of the separated lipid classes (detection limits of approximately 25–50 ng per lipid class) [46] even if differences in the fatty acyl compositions have a significant impact on the results. The mechanism behind this is quite complex: while the sulfur in sulfuric acid is reduced from +VI to +IV, the carbons of the lipid are partially oxidized to elementary carbon (-II (CH2) ⟶ ±0 (C)) and some lipid is completely converted into CO2 (-II (CH2) ⟶ +IV (CO2)) [47].
Many other reagents, such as potassium dichromate (5%), copper sulfate in 40% H2SO4 or a 3–6% solution of cupric acetate in 8–10% phosphoric acid (H3PO4) [48] were also indicated to represent useful stains. Readers interested in a survey of such destructive approaches are referred to [49] and the references cited therein. A survey is also available in [50]. One cautionary note seems also appropriate: TLC plates with polymer (plastic) layers are normally not suitable for these assays, as the polymer is depleted by the corrosive agents. Glass plates should be used instead.
Some selected destructive staining approaches are summarized in Table 3:

3. Spectroscopic Detection of Lipids on the TLC Plate

Although there have been numerous attempts to use spectroscopic methods to elucidate details of the detected analytes directly on a TLC plate, most techniques (except for UV spectroscopy) have not achieved significant practical relevance.

3.1. UV Spectroscopy and/or Quenching

Detection using ultraviolet (UV) light, and, to a more limited extent, fluorescence, is normally not possible with standard lipids. However, it can be applied to oxidized lipids (e.g., diene conjugates) [58] as well as to some artificial lipids containing a fluorophore. This can be easily performed by examining dried TLC plates under both 254 and 366 nm UV light. The comparison of both wavelengths enables conclusions about the compounds of interest [59].
The quenching of TLC plate fluorescence is another convenient method if plates with a fluorescent indicator (F254) are used. If these dried plates are examined under 254 nm UV light, compounds absorbing this wavelength appear as dark spots against a green or white fluorescence background. It is important to note that the presence of the fluorescent indicator may affect the migration behavior of lipids on TLC plates [60] since the silica gel is slightly altered by the fluorophore.
The applications of other spectroscopic methods are normally hampered by the following aspects:
(1)
Methods such as infrared (IR) or RAMAN spectroscopy lack specificity, i.e., these methods only detect some characteristic functional groups that enable the detection of lipids (e.g., the carbonyl groups) but without providing further details [61]. Nevertheless, the application of IR in combination with matrix-assisted laser desorption/ionization (MALDI) MS has recently been described [62].
(2)
Nuclear magnetic resonance (NMR)-based methods are powerful tools enabling structural as well as quantitative analysis. However, the sensitivity is very low and, accordingly, NMR is only scarcely used in the context of TLC [63] and, to the best of our knowledge, has never been used for the characterization of lipids on a TLC plate.
(3)
MS is widely used to elucidate the lipid fractions on a TLC plate [64]. In the following sections, we will focus on the coupling between TLC and soft ionization MS (electrospray ionization (ESI) and MALDI), discussing the application of extraction-based and desorption-based techniques separately.

3.2. Extraction-Based Analysis of Stained TLC Plates

One of the most widely used MS techniques in the lipid field is ESI MS. Here, the sample is introduced into the mass spectrometer as an analyte solution in a polar, ESI-compatible solvent such as methanol. Even though the extraction efficiency of chloroform would be higher, chloroform is not a suitable solvent for ESI MS due to its apolarity and the low ion yields of the analytes which would be obtained under these conditions [65]. Additionally, unwanted dichlorocarbene generation may occur [66] and due to its aggressiveness, chloroform harms capillaries and all plastic-containing parts of the machinery.
ESI and other soft-ion ionization MS techniques (as well as MALDI and atmospheric pressure chemical ionization (APCI)) are characterized by a considerable ion suppression effect, meaning that different analyte species are not detected according to their concentration but their tendency to yield ions in the gas phase [67]. Overcoming the ion suppression effect is the most important reason why ESI MS is normally coupled with liquid chromatography (LC) to separate the different lipid classes (e.g., according to their headgroups) prior to MS analysis.
When TLC is used instead of LC, a slightly different analytical approach is employed [68]: as soon as the lipid mixture is separated on the TLC plate, the obtained individual lipid spots (monitored by any non-destructive dye such as primuline, which does not alter the molecular weight of the lipids) are extracted and the resulting analyte solution is directly transferred from the TLC plate to the mass spectrometer through a dedicated TLC-MS interface. At least two of such devices are commercially available from CAMAG (TLC-MS Interface ®) and Advion (Plate Express ®).
Coupling ESI with TLC is simple to establish and useful, making it suitable for many applications. A review covering selected lipid applications is available in [69]. A schematic survey which illustrates the typical workflow is shown in Figure 2.
The available devices operate in a very similar way: the oval or circular elution head is positioned directly on the TLC spot of interest, which was previously stained by a non-destructive stain such as primuline and marked with a pencil under UV light. An ESI-compatible solvent is subsequently pumped through a dedicated elution head. The lipid within the spot is extracted via an inlet capillary. Any potential silica gel contaminants are filtered out, and the eluate is directed into the mass spectrometer via the outlet capillary. This method was introduced by Heinrich Luftmann about 20 years ago [70]. A key advantage of this approach is that each type of MS, using different ionization techniques and mass analyzers, could be used. However, typically, ESI mass spectrometers are used—presumably due to their broad applicability (they detect most lipids with reservations regarding very apolar species such as cholesteryl esters).
Although this coupling is a powerful method, there is a limitation: due to the small solvent volumes which are typically used, the time for acquisition is short and, thus, the time to record MS/MS spectra is limited. Fortunately, this is mainly a serious problem in the case of completely unknown lipid classes.

4. Desorption Techniques

Compared to elution-based ionization techniques [71], methods using desorption-based ionization, for example, desorption electrospray ionization (DESI) [72] and MALDI, allow lipid detection without the re-elution of the corresponding TLC spot. These methods are comprehensively discussed in [73], where a survey of densitometric methods is also provided. In this review, we will focus on MALDI MS, as this technique is the primary one used in our laboratory [74].
Independent of the used MS method, desorption-based techniques offer some advantages in comparison to extraction-based techniques:
(1)
There is no need to extract the sample from the TLC plate, since the MS analysis is performed directly on the TLC plate. This is convenient and bears no risk of analyte loss upon the extraction process. However, it remains unclear whether all analytes are desorbed to the same extent from the TLC plate.
(2)
Since the analysis of a dedicated spot can be performed within a few seconds, there is no risk of sample alteration, such as unwanted lipid (per)oxidation. This particularly applies because the TLC plate is under high vacuum during data acquisition. For instance, we successfully analyzed lipids in sperm, which are rich in docosahexaenoic acid with six double bonds, without a significant sign of oxidation [75].
(3)
Since the spatial resolution is determined by the MALDI laser spot size, the resolving power is much higher compared to common staining and visual inspection [74]. Thus, TLC/MALDI is able to resolve different components within a single TLC spot that cannot be resolved by staining alone, for example, phospholipid species with the same headgroup but different fatty acyl residues.
Unfortunately, so far there have been almost no investigations comparing densitometry, extraction-based methods and desorption MS. Krüger and coworkers [76] compared compositional details of soybean and sunflower lecithins (PCs) using HPTLC with fluorescence detection and ESI as well as MALDI MS evaluation. Similar data were obtained with all three methods, although there was a slight, but remarkable mass shift in the MALDI mass spectra. The authors explained the shift by slight differences in the height variations (“roughness”) of the TLC surface caused by cutting a larger TLC plate into smaller ones. This effect, however, can be overcome by using industrially manufactured, smaller TLC plates which are now also available in high-purity (MS-grade) versions [77].

5. Summary

Thin-layer chromatography represents unequivocally a powerful method to perform compositional studies of lipid mixtures, although the accuracy and sensitivity of modern MS-based lipidomics studies can be hardly achieved.
Traditionally, the developed (typically normal phase) TLC plates are stained with suitable dyes and the obtained lipid fractions subsequently quantified by densitometry. This paper discussed numerous dyes differing in their specificity. Many of them are commercially available. A short summary of the data within this review is shown in Figure 3.
Although there are a lot of references available, it has to be admitted that the precise mechanisms by which some dyes interact with lipids are still not fully understood. Therefore, most dyes are used based on their availability and/or whether they were established in a particular laboratory. Regarding dye selection, the optimal method depends on whether destructive analysis and sensitivity are prioritized (e.g., cupric sulfate or phosphomolybdic acid staining with charring) or if non-destructive detection is needed (e.g., primuline–UV). Generally, destructive approaches are normally slightly more sensitive [23]. This review contributes additional guidance for selecting the most suitable dye.
The lipid class is often already obvious from the Rf ratio of known reference compounds, but the evaluation of the fatty acyl composition of the different (phospho)lipids is much more challenging and requires more sophisticated approaches such as TLC combined with MS. Even though some further methodological developments/improvements are needed, there is clear evidence that this coupling approach will further promote TLC as a powerful, versatile analytical tool with advantages in comparison to LC.
Furthermore, there are already established workflows in which the headgroup of a lipid is determined by TLC first and afterwards the spot of interest is hydrolyzed and the released free fatty acids are determined by GC [78].

Author Contributions

J.W.S., J.L. and K.M.E. contributed equally. The conceptualization was made by J.W.S., J.L. and K.M.E. The original draft was written by J.W.S., J.L. and K.M.E. J.S. and K.M.E. were responsible for the dyes, while the spectroscopic part was mainly written by K.M.E. The final editing as well as funding acquisition was performed by J.S. All authors have read and agreed to the published version of the manuscript.

Funding

The authors were supported by the German Research Foundation (EN 1279/3-1 (511424302) to KME; SCHI 476/26-1 (531487925) to JS).

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
APCIatmospheric pressure chemical ionization
DESIdesorption electrospray ionization
ESIelectrospray ionization
F254fluorescent indicator
GCgas chromatography
HPLChigh-performance liquid chromatography
HPTLChigh-performance thin-layer chromatography
IRinfrared
LCliquid chromatography
LPClysophosphatidylcholine
MALDImatrix-assisted laser desorption/ionization
MSmass spectrometry
MS/MStandem mass spectrometry
NMRnuclear magnetic resonance
NPnormal phase
PBH1-pyrenebutyric hydrazide
PCphosphatidylcholine
PEphosphatidylethanolamine
PLphospholipid
PSphosphatidylserine
RPreversed phase
SMsphingomyelin
TLCthin-layer chromatography
UVultraviolet

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Lipidology 03 00003 g001
Figure 1. One selected phospholipid (PL) (a phosphatidylcholine (PC)) with its different functional groups is shown in the upper part of the figure. Fatty acyl chains are shown in blue, double bonds are highlighted in green and the phospholipid headgroup is depicted in red. The different groups which are normally monitored during the staining process are emphasized in the lower part of the figure. The acyl residues can be monitored by charring while the double bonds are stained by iodine (I2). More specifically, the phosphate is determined by the Dittmer–Lester agent and the ammonia group by the Dragendorff reagent. Mass spectrometry (MS) combined with tandem MS (MS/MS) enables the detection of the entire molecule as well as (at least) the characteristic headgroup and does not require staining. Thus, this is probably the most powerful approach.
Figure 1. One selected phospholipid (PL) (a phosphatidylcholine (PC)) with its different functional groups is shown in the upper part of the figure. Fatty acyl chains are shown in blue, double bonds are highlighted in green and the phospholipid headgroup is depicted in red. The different groups which are normally monitored during the staining process are emphasized in the lower part of the figure. The acyl residues can be monitored by charring while the double bonds are stained by iodine (I2). More specifically, the phosphate is determined by the Dittmer–Lester agent and the ammonia group by the Dragendorff reagent. Mass spectrometry (MS) combined with tandem MS (MS/MS) enables the detection of the entire molecule as well as (at least) the characteristic headgroup and does not require staining. Thus, this is probably the most powerful approach.
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Figure 2. Schematic workflow of TLC/MS coupling: the lipid mixture is separated by conventional TLC. Afterwards, the separated lipids are stained by any non-destructive dye (for instance, primuline), automatically re-eluted and infused into the mass spectrometer. Reprinted with permission from Elsevier [67].
Figure 2. Schematic workflow of TLC/MS coupling: the lipid mixture is separated by conventional TLC. Afterwards, the separated lipids are stained by any non-destructive dye (for instance, primuline), automatically re-eluted and infused into the mass spectrometer. Reprinted with permission from Elsevier [67].
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Figure 3. Survey of the different (staining) methods which can be used to obtain structural information from the spots on a developed TLC plate. We recommend (in red) the use of primuline staining if a classical staining approach has to be applied. Although more expensive, ESI MS offers the maximum amount of information and is being increasingly used.
Figure 3. Survey of the different (staining) methods which can be used to obtain structural information from the spots on a developed TLC plate. We recommend (in red) the use of primuline staining if a classical staining approach has to be applied. Although more expensive, ESI MS offers the maximum amount of information and is being increasingly used.
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Table 2. Selected staining approaches for lipids separated on TLC plates.
Table 2. Selected staining approaches for lipids separated on TLC plates.
MethodCharacteristics/SpecificityReference
Iodine (I2)I2 is non-covalently bound to lipids; spot intensity depends on the number of double bonds. Rather inexpensive and simple [33].[27]
Rhodamine 6GFor the detection of lipids at long-wavelength UV. Suggested to represent a “lipidomics” approach because virtually all lipids are detectable [34].[35]
Coomassie brilliant blueStains virtually all lipid classes. The sensitivity is about 0.05 to 0.5 µg per lipid. Was recently combined with ESI MS detection of the re-eluted lipids [32].[31]
Nile red (fluorescent)Detects most lipids, although the fluorescence intensity varies among the lipid classes. Also, staining is stronger for unsaturated lipids than for saturated ones. To the best of our knowledge, it is the most sensitive assay. Detection limit of 25–100 ng. [36]
HematoporphyrinFluorimetric detection of low nanogram amounts of cholesterol, cholesteryl esters, triolein, sphingomyelin and lecithin by using an argon laser (488 nm). Copper-(II)-nitrate solution attenuates the background fluorescence.[37]
BerberineSuitable for the estimation of the length of the alkyl chains since the fluorescent emission increases in the presence of molecules with long hydrocarbon chains.[38]
2′,7′-dichloro-fluoresceinFor lipid samples containing 2.5 up to 20 μg lipid. 2′,7′-dichlorofluorescein detects all lipid classes while sulfuric acid/ethanol spray reagent (used for means of comparison) could not detect saturated fatty acids and LPC.[39,40]
Primuline (Direct yellow 59)Non-covalent interaction with lipids. Perfectly suited for combination with MS detection because primuline can be quantitatively removed from the lipid by exposition to vacuum and/or changing the polarity of the solvent.[41]
Table 3. Destructive staining approaches for lipids.
Table 3. Destructive staining approaches for lipids.
MethodCharacteristicsRef.
Phosphomolybdic acid (MDA)
(for detection of reducing compounds such as cholesterol and selected bile acids)		Charring until lipids are visible as brown, gray, or black spots. MDA can be used for NP and RP plates. In the case of octadecylsilica materials, however, the MDA also reacts with some of the C18 groups. The consequence of this is a dark background and, therefore, poorly sensitive analyte detection.[51]
Phosphotungstic acidFor the detection of cholesterol and its esters, reducing compounds, lipids, sterols and steroids. Cholesterol esters will produce red spots. Significantly less often used than MDA but often applied in tissue analysis.[52]
Potassium dichromate (in concentrated sulfuric acid)Brown, gray, or black spots result. Similar to MDA.[53]
Potassium permanganate/sulfuric acidDo not use this reagent on polymer-bound TLC plates since it will destroy the binder; use it only with G (gyp-sum) binder plates! Excellent for functional groups sensitive to oxidation. Alkenes and alkynes will appear readily on a TLC plate following immersion into the stain as bright yellow spots on a bright purple background. Alcohols, amines, sulfides, thiols and other oxidizable functional groups often require slight heating.[54]
Copper (II) sulfate in phosphoric acidOften considered as the optimum method of lipid staining since it stains all major lipid classes to an identical extent. Thus, differences in spot intensities reflect different concentrations. Fast (dipping for 3 s is sufficient). About 1.7 to 2.0 μg of lipids are detectable [55].[56]
Ceric ammonium molybdate (CAM) staining with charringRemarkable sensitivity of just about 130 ng of lipid. Useful if the amount of lipids is limited.[22]
Orcinol or resorcinol in sulfuric acidSpots containing glycolipids develop a pink-violet color on a white background. Detection limit of approximately 0.5 nmol.[57]
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Schubarth, J.W.; Leopold, J.; Engel, K.M.; Schiller, J. Lipid Analysis by Thin-Layer Chromatography—Detection, Staining and Derivatization. Lipidology 2026, 3, 3. https://doi.org/10.3390/lipidology3010003

AMA Style

Schubarth JW, Leopold J, Engel KM, Schiller J. Lipid Analysis by Thin-Layer Chromatography—Detection, Staining and Derivatization. Lipidology. 2026; 3(1):3. https://doi.org/10.3390/lipidology3010003

Chicago/Turabian Style

Schubarth, Johanna W., Jenny Leopold, Kathrin M. Engel, and Jürgen Schiller. 2026. "Lipid Analysis by Thin-Layer Chromatography—Detection, Staining and Derivatization" Lipidology 3, no. 1: 3. https://doi.org/10.3390/lipidology3010003

APA Style

Schubarth, J. W., Leopold, J., Engel, K. M., & Schiller, J. (2026). Lipid Analysis by Thin-Layer Chromatography—Detection, Staining and Derivatization. Lipidology, 3(1), 3. https://doi.org/10.3390/lipidology3010003

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