Pandohee Et Al-2015-Journal of Separation Science

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12 15

JOURNAL OF

SEPARATIONSCIENCE

ISSN 1615-9306 · JSSCCJ 38 (12) 2007–2192 (2015) · Vol. 38 · No. 12 · June 2015 · D 10609

JSS

MethodsChromatography · Electroseparation

ApplicationsBiomedicine · Foods · Environment

2024 J. Sep. Sci. 2015, 38, 2024–2032

Jessica Pandohee1

Brendan J. Holland2

Bingshan Li5Takuya Tsuzuki3Paul G. Stevenson2

Neil W. Barnett2

James R. Pearson4

Oliver A.H. Jones1

Xavier A. Conlan2

1School of Applied Sciences,RMIT University, Melbourne,Victoria, Australia

2Centre for Chemistry andBiotechnology, School of Lifeand Environmental Sciences,Deakin University, Geelong,Victoria, Australia

3Research School of Engineering,Australian National University,Canberra, Australia

4Victoria Police Forensic ServicesDepartment, Macleod, Victoria,Australia

5Institute for Frontier Materials,Deakin University, Geelong,Victoria, Australia

Received January 28, 2015Revised March 25, 2015Accepted March 27, 2015

Research Article

Screening of cannabinoids inindustrial-grade hemp usingtwo-dimensional liquid chromatographycoupled with acidic potassiumpermanganate chemiluminescence detection

Widely known for its recreational use, the cannabis plant also has the potential to act asan antibacterial agent in the medicinal field. The analysis of cannabis plants/products inboth pharmacological and forensic studies often requires the separation of compounds ofinterest and/or accurate identification of the whole cannabinoid profile. In order to pro-vide a complete separation and detection of cannabinoids, a new two-dimensional liquidchromatography method has been developed using acidic potassium permanganate chemi-luminescence detection, which has been shown to be selective for cannabinoids. This wascarried out using a Luna 100 Å CN column and a Poroshell 120 EC-C18 column in thefirst and second dimensions, respectively. The method has utilized a large amount of theavailable separation space with a spreading angle of 48.4� and a correlation of 0.66 allowingthe determination of more than 120 constituents and mass spectral identification of tencannabinoids in a single analytical run. The method has the potential to improve researchinvolved in the characterization of sensitive, complex matrices.

Keywords: Cannabis / Offline chromatography / Profiling / 2D chromatographyDOI 10.1002/jssc.201500088

� Additional supporting information may be found in the online version of this articleat the publisher’s web-site

1 Introduction

Comprising over 525 compounds, cannabis is a complex plantthat includes three different species (Cannabis sativa, C. in-dica, and C. ruderalis) that has been of interest for its biologi-cally active constituents for hundreds of years [1, 2]. It is wellknown as a recreational drug due to its psychoactive effect, at-tributed to the cannabinoid �9-tetrahydrocannabinol (THC)[3]. Other cannabinoids of importance are cannabidiol (CBD),cannabichromene (CBC), cannabigerol (CBG), and cannabi-nol (CBN), all of which have been shown to possess potent ac-tivity against a variety of methicillin-resistant bacterial strainsof current clinical relevance [4, 5]. Many other cannabinoidshave also been identified which have been associated with arange of potential medicinal uses such as anti-inflammatory,antibiotic, antifungal, analgesic, and antioxidant compounds

Correspondence: Dr. Xavier A. Conlan, Centre for Chemistryand Biotechnology, School of Life and Environmental Sciences,Deakin University, Pigdons Road, Geelong, Victoria, AustraliaE-mail: [email protected]

Abbreviations: CBC, cannabichromene; CBD, cannabidiol;CBGA, cannabigerolic acid; CBG, cannabigerol; CBN, cann-abinol; THC, �9-tetrahydrocannabinol

[6]. However, the use of cannabis as a medicine is still dis-puted because of socio-political pressure [1].

Hillig and Mahlberg [7] classified the cannabis plantsinto three chemotypes depending on the THC/CBD ratio;(i) drug-type plants with a high THC/CBD ratio (>>1:1),(ii) intermediate type plants with a THC/CBD ratio closeto 1:1, and (iii) fiber-type plants with low THC/CBD ratios(<<1:1). The latter form, also known as hemp is grown forits seeds and fibers for agricultural and industrial purposes.Hemp is known to be some of the best and most durablefibers of natural origin and has been used to make paper,banknotes, ropes, and sails for ships [1]. Hemp seeds are alsonutritious containing a high oil content, namely linoleic acid,omega-3, and omega-6, essential fatty acids, and proteins;they may be consumed in their natural form or as an oil, aseed milk, or as a herbal infusion [8, 9].

Considering the complex nature and uses of thecannabis/hemp plant, the need for separating its constituentsfor further research is evident, but the degree of separation re-quired varies with the purpose of the next study. For example,forensic analyses for the identification/classification of fiberor drug-type, and source tracing demand an accurate finger-print of the entire cannabinoid profile [2]. In contrast, phar-macology often requires the isolation of a pure constituent(s)of the plant extract [2]. Consequently, improvement in the

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J. Sep. Sci. 2015, 38, 2024–2032 Liquid Chromatography 2025

separation of such cannabinoids is essential to achieve base-line resolution for both isolation and profiling of the samples.

Currently, GC is the method of choice for creatingcannabis profiles and chemical fingerprints for attributingthe country of origin or conditions of cultivation, as it allowsidentification of a large variety of cannabinoids with very highresolution, particularly when coupled with MS [2]. However,the high temperatures required for sample vaporization be-fore injection causes the decarboxylation of the acidic formsof cannabinoids and thermal degradation of the sample, thuspreventing analysis of cannabis in its natural state [2]. Further-more, this thermal conversion of acidic cannabinoids seemsto be incomplete, resulting in a nonrepresentative analysis ofthe sample [10]. The quantification of cannabinoids by GCtherefore requires a time-consuming derivatization beforeanalysis [10].

HPLC allows the simultaneous detection of both acidicand neutral cannabinoids with no need of derivatization [11].However, the majority of HPLC methods for cannabis analy-sis described in the literature either failed to separate all thecannabinoids present and/or were not validated according toappropriate guidelines [12]. The complex composition of thematerial means that the separation of major cannabinoids isnot easily achieved and significant peak overlap occurs be-tween CBD/CBG and CBN/cannabigerolic acid (CBGA) [5].The use of MS coupled to HPLC may be a solution to re-solve cannabinoids of interest in a single analytical run. Thismethod has been previously used to separate acidic and neu-tral cannabinoids allowing selective identification in hashishsamples with no derivatization or peak collection [13] anddetermination of up to five cannabinoids in hemp [14]. Nev-ertheless, LC–MS does not allow characterization of an entirecannabis sample, but the determination of specific analytes.However, for characterization and QC purposes, determina-tion of cannabinoids in only one single run with high selec-tivity would be advantageous [14].

It is evident that 1D chromatographic techniques havesignificant limitations, especially when chromatographic res-olution of numerous compounds is desired [15]. A gain inseparation power/peak capacity can be achieved using 2Dchromatography [16–18]. The technique involves combiningtwo dimensions of different separation mechanisms in se-ries. Fractions from the first dimension are collected andre-injected into the second dimension where they are fur-ther separated by an orthogonal (different) separation mech-anism [16]. This provides a gain in peak capacity as the totalpeak capacity of the 2D system is the product of the peakcapacity of each dimension [16].

The 2D LC–UV/chemiluminescence (2DLC–UV/CL)methodology described here was developed to separate thecannabinoids in industrial-grade cannabis efficiently. The or-thogonal separation was then used to compare the chemicalprofiles of samples of cannabis bark, hurd, and leaves ofboth sexes of the plant. The acidic potassium permanganatechemiluminescence detection system, provided an indicationof the chemical activity of compounds in the cannabis sam-ples [19]. To the best of the authors’ knowledge, this is the

first comprehensive 2D HPLC study of cannabis and has highpotential for future research.

2 Materials and methods

2.1 Chemicals

All chemicals were analytical grade reagents, unless other-wise stated. Potassium permanganate, ethyl acetate, sodiumsulfate (anhydrous powder) were obtained from Chem-Supply (Gillman SA, Australia). Sodium polyphosphate(crystals, +80 mesh, 96%), sodium thiosulfate (powder)were purchased from Sigma-Aldrich (Castle Hill, NSW,Australia). Methanol (Isocratic, HPLC grade) was acquiredfrom Scharlau Chemie (Gillman SA, Australia). Sulfuric acid(�98.0%) was sourced from Merck (Kilsyth, VIC, Australia).

2.2 Extraction of cannabinoids from hemp

Industrial-grade hemp samples, root hurd and leaf (Com-mins Stainless Manufacturing Farm, Whitton, NSW,Australia) were washed under reverse osmosis water(18.2 M�), dried at 60�C, ground into a powder, and passedthrough a 368 �m screen. Samples were prepared from 0.5 gof the powder by solvent extraction with 300 mL of ethyl ac-etate for 1.5 h at 78�C using a Soxhlet apparatus. The extractwas removed and the extraction repeated with another 300 mLof solvent for 1 h. The two extracts were combined, evaporatedto dryness under vacuum at 40�C, then reconstituted in ethylacetate (20 mL) and stored at <4�C until use.

2.3 Preparation of acidic potassium permanganate

chemiluminescence reagent

The acidic potassium permanganate solution was prepared bydissolving sodium polyphosphate (10 g/L) in distilled water,adding potassium permanganate (1.9 mM) and adjusting thepH to 2.50 with concentrated sulfuric acid followed by theaddition of sodium thiosulfate (1.0 mM) from a 0.1 M stocksolution.

2.4 Chromatography columns

The following columns were used for selectivity studies:Eclipse XDB-C18 (Agilent, 4.60 mm × 150 mm, 5 �m parti-cle diameter, Pd), Luna 5u CN 100Å (Phenomenex, 4.60 mm× 150 mm, 5 �m Pd), Luna 5u HILIC 200Å (Phenomenex,4.60 mm × 150 mm, 5 �m Pd, Luna 5u NH2 100Å (Phe-nomenex, 4.60 mm × 150 mm, 5 �m Pd), Poroshell 120EC-C18 (Agilent, 4.60 mm × 50 mm, 2.7 �m Pd), ChromolithRP18e (Merck, 4.60 mm × 100 mm, 5 �m Pd).

Based on the data obtained from this study (discussed inSections 3.1 and 3.2), the Luna 5u CN 100Å and Poroshell


2026 J. Pandohee et al. J. Sep. Sci. 2015, 38, 2024–2032

Figure 1. Uni-dimensional separation of female leaf ethyl acetate extract on (A) HILIC, (B) NH2, (C) CN, and (D) C18 showing selectivity ofeach stationary phase toward the cannabinoids.

120 EC-C18 were used as first and second dimensions, re-spectively for the 2D analyses (see Section 2.7 for methoddetails).

2.5 Instrumentation

Chromatographic analysis was carried out on a 1200 Se-ries HPLC system (Agilent Technologies, Mulgrave, VIC,Australia), equipped with two quaternary pumps, solventdegassers, an autosampler, and a continuously variable-wavelength detector at 220 nm, followed by chemilumi-nescence detection. The two columns, a Luna 100 Å CN(Phenomenex, 4.60 mm × 150 mm, 5 �m Pd) for the firstdimension and Poroshell 120 EC-C18 (Agilent, 4.60 mm ×50 mm, 2.7 �m Pd) for the second dimension were con-nected via an electronically controlled two-position ten-portvalve fitted with microelectric two-position valve actuatorsthat allowed alternate sampling of the eluent from the first di-mension into the second dimension. Before use in the HPLCsystem, all sample solutions and solvents (except HPLC-gradesolvents) were filtered through a 0.45 �m nylon membrane.

2.7 2D configuration

Data of the selectivity study led to a linear gradient conditionbeing employed on both columns, starting from an initial mo-bile phase composition of 30% methanol, running to a final

mobile phase composition of 100% methanol at a rate of 8.6%per min and initial mobile phase composition of 100% water,running to a final phase composition of 100% methanol at arate of 33% per min for the first and second columns, respec-tively. The injection size was 100 �L in the first dimensionand a flow rate of 1 mL/min was used for both dimensions.The 2D-HPLC analysis involved transferring the eluent flowstream from the first dimension (200 �L) into the seconddimension (using a heart-cutting process) after the seconddimension separation was completed a second sample wasinjected into the first dimension. The heart-cutting processwas repeated, each time sampling every second 200 �L por-tion of the eluent from the first dimension 33 times, thisresulted in a total analysis time of 12 h.

Postcolumn acidic potassium permanganate chemilumi-nescence was generated using the custom-built manifoldpreviously described [20]. The chemiluminescence reagent,propelled at a flow rate of 1.2 mL/min merged with the HPLCeluent at a T-piece junction and the light emitted from the re-acting mixture was detected with a custom-built flow-throughluminometer, which comprised a coiled flow cell, mountedflush against the window of an electron tube photomultipliertube (model 9828SB, ETP) set at a constant voltage of 900 Vfrom a stable power supply (PM20D, ETP) by a voltage di-vider (C611, ETP). An analog to digital interface box (AgilentTechnologies) was used to convert the signal from the chemi-luminescence detector. 2D data plotting and peak pickingwere undertaken using custom written code in Mathematica(version 8, Wolfram Research, Champaign, IL, USA).



Figure 2. Optimized uni-dimensional separations of female leaf extract on (A) CN column with vertical gridlines representing fraction cuts(B) Poroshell EC-C18.

2.8 MS

An HPLC, electrospray time-of-flight (HPLC–ESI-TOF) massspectrometer (Agilent 6210 MSD, Agilent Technologies) wasused in a positive mode to identify the cannabinoids under thefollowing conditions: drying gas: nitrogen (7 mL/min, 350�C);nebulizer gas: nitrogen (16 psi); capillary voltage 4.0 kV; va-porizer temperature 350�C, and cone voltage 60 V. An injec-tion volume of 10 �L was used. Reference masses used werethe ESI tuning mix G2412A (Agilent Technologies).

3 Results and discussion

3.1 Selectivity testing – First dimension method

development

Selectivity testing was performed using HILIC, amino (NH2),cyano (CN), and C18 columns to determine which stationaryphase was most suited for the analysis. (Fig. 1 panels A–D).The HILIC column (Fig. 1A displayed a limited number ofresolved peaks and the majority of the components eluted by8 min. The NH2 column (Fig. 1B) utilized a large amount ofseparation space; however, there were issues with the broadpeaks observed in particular with the variations in the noisein the detection system. Nevertheless, judgements on the col-umn efficiency could still be made. The CN column (Fig. 1C)performed similarly to the NH2 column in terms of use ofseparation space but better resolution of the components ispresent (this takes into account the signal-to-noise differencesbetween these two runs). Separation was found to be im-proved by utilizing the unused space from 3 to 7 min. TheC18 (d) displayed the best use of separation.

Based on these findings, the CN column was deemed tooffer the best separation for the first dimension separationthat would be coupled with the C18. Nonetheless, to use theseparation space of the CN column efficiently and reduce theseparation time, a series of additional tests were undertaken.First, each consecutive separation involved an increment of10% methanol in the initial mobile phase composition. The

initial mobile phase content range investigated was from 0 to70% methanol over a gradient of 12 min followed by 3 min of100% methanol. The separation starting with 30% methanol(at a rate of 5.8% per min) was found to be the most ap-propriate in gaining in usage of the separation space whilesimultaneously maintaining a fair resolution. Then, a seriesof experiments with increasing speed of the previously op-timized gradient was performed with their separation timebeing reduced from 12 to 6 min Electronic Supporting Infor-mation (ESI1) It is important to note that at some point thetrade-off between separation effectiveness and speed of theanalysis needs to be carried out. In this case, the separationwith a gradient increasing at a rate of 8.75% methanol perminute was chosen as the first dimension protocol as it of-fered a reasonable separation in a practical length of time, theminor loss in resolution at this point is trivial since furtherseparations on the second dimension allowed a significantgain in peak capacity.

3.2 Second dimension method development

A comprehensive offline approach, where the first dimensionis repeatedly performed for fraction collection and followedby subsequent injection in the second dimension, allows highseparation power to be achieved [21]. However, a very longanalysis time is a cost for the substantial gain in peak capacity.Consequently, a series of C18 columns was tested to identifythe best type of column for this separation (considering bothpeak capacity and time constraint).

The performance of a Poroshell EC-C18 and a mono-lithic column were compared using a linear gradient at aflow rate of 1 mL/min, starting with 0% methanol to 100%methanol in 3 min followed by 3 min of 100% methanol. Thechromatograms obtained are shown in (ESI1). The PoroshellEC-C18 was found to have a shorter analysis time and moreimportantly a better resolution power than the monolith. Anadvantage on the short column is that it provides faster anal-yses and in this case the resolution power was noteworthycompared to the monolith. Further separations were carried



Figure 3. (A) 2D plot showing separation space used in a male leaf ethyl acetate separation (B) Geometric approach to factor analysisresults of five cannabis samples.

out to reduce the analysis time on the C18 but a negligiblegain in the usage of the separation was achieved. Therefore,the original method, starting from 0% methanol to 100%methanol, was chosen for the 2D-HPLC runs.

After the columns had been chosen from the selectivitystudy, each separation was studied to ensure the columnscould be coupled in the 2D system in the most effective man-ner. The stationary phase is the most important considerationof any separation, followed by the mobile phase composi-tion and the effect it may have on separation selectivity. Thisconsideration is most important for sample transfer modula-tion to suppress band broadening and achieve peak focusing,peak capacity, orthogonality, and sampling frequency [22].The main challenge remains to maintain efficient coupling ofthe columns while preserving the mobile phase/column com-patibility. Limited solvent miscibility and/or solvent strengthproblems may significantly affect the retention and selectivitybut also peak shapes and band broadening in the second di-mension. When coupling two dimensions for 2D-HPLC anal-yses, the use of two completely different column-retentionmechanisms is highly beneficial for a gain in orthogonality,hence maximizing the increase in peak capacity [16]. How-ever, a major issue faced by several research groups [23, 24]is solvent incompatibility of the nonpolar normal-phase sol-vents and the polar reversed-phase solvents. To remedy tothis, the same solvents (water/methanol gradient) were usedin both columns as previously described [25, 26].

In Fig. 2, the optimized separations of the CN first dimen-sion (A) and the C18 second dimension (B) can be observed.The chromatogram generated on the CN column shows thecomplexity of the hemp extract and that accurate chemical

characterization of the sample cannot be obtained from theanalysis at this stage. Moreover, it is evident that the peakcapacity has been exceeded. The vertical gridlines on ESI1represent the time at which fractions were collected and in-jected in the C18 second dimension. Although the C18 wasselective and showed some degree of resolution, the contin-uum of sample from 3 to 5 min shows that the peak capacityhas again been exceeded. ESI2 illustrates a heart cut fractionfrom the first dimension at 2.2 min of female (A) and male(B) leaves. It is very interesting to observe the strong similar-ity between both separations with the peak at 3 min as thesole difference. These fraction-cuts also show the number ofunresolved peaks from the first dimension and the need for2D-HPLC.

To assess the separation space occupied by the samplecomponents and orthogonality of the system [27, 28], a ge-ometric approach to factor analysis (GAFA) was carried outon each of the five cannabis sample. The 2D plot in Fig. 3shows that the separation partially occupies the availablespace, which is indicated by the spreading angle of 48.4�.The larger the spreading angles the more separation spacewill be used by the sample components and a spreading an-gle of 90� would indicate full usage of the separation space.While most spreading angles obtained for the cannabis sepa-rations were < 50�, correlation coefficients also indicated thatthe dimensions were correlated with a high correlation coef-ficient of up to 0.80 with the female leaf extract, which wouldprobably be explained by the use of RP separations in bothdimensions. It is of note that achieving a separation that usesthe 2D space effectively can be very challenging and involvesthe usage of columns that are as dissimilar as possible. The



Figure 4. (A) Typical 2D plot of hemp with identified cannabinoids a-CBV b-(CBCV,CBDV,CBLV), c-CBGV, d-CBN, e-(CBC,CBD, CBL), f-CBG,g-CBE, h-CBT, i-CBNA, j-(CBCA,CBDA,CBLA), k-CBGA.

CN column comprises nitrile groups bonded to silica-basedcolumn and has a high affinity for polar compounds. The C18

is a superficially porous micro-particulate column packingwith a solid silica core and a porous silica outer layer. Unlikethe CN column, the C18 reduces strong adsorption of basicand highly polar compounds and these were the two mostdiffering column chemistries.

3.3 Tentative identification of the cannabinoids

using LC-MS

The known masses of the eight neutral cannabinoid classes,together with that of the varin (cannabinoids with a propylgroup) and acidic cannabinoids (Fig. 3) were used to obtainextracted ion chromatograms (EIC) from each sample. Theretention times of the assigned cannabinoids’ peaks from theEIC of both the CN and short C18 chromatograms were sub-sequently determined and utilized as coordinates to estimatethe retention position of the cannabinoids in the 2D contourplots, as shown in Fig. 4. The overall separation has beendivided in three regions: region 1 containing compounds nothighly retained on the CN column, region 2 containing com-pounds highly retained by the Poroshell EC-C18, and region3 compounds highly retained on both columns. Cannabi-noids of interest were found to elute in region 2. AlthoughTHC, CBC, CBD, and CBL have the same molecular weightof 314.46 amu, identification of each component was made

with the support of previous LC-MS results [29].The sam-ple being fiber-type cannabis would contain CBD as its mostabundant cannabinoid (assigned to peak with highest inten-sity) and the presence of THC would be negligible. CBC wasshown to elute after CBD, so it was assigned to the peak witha retention time of 5 min, leaving CBL as the first peak of thered EIC. It is not surprising to see a higher content of CBLcompared to CBC, as the latter degrades into CBL indicatingthat the sample may have undergone some degradation. Thiscan also be established by the presence of only two of theacid cannabinoids (CBCA/CBDA/CBLA). It is evident thatCBD has a major role in the antibacterial properties of hemp;the presence of other cannabinoids such as CBL, CBC, andperhaps CBGA and CBG suggests synergistic activity.

One of the key outcomes of this technology is the abil-ity to generate in-depth chemical fingerprints of the complexhemp extracts, the power of this can be observed through the2D chromatograms of extracts from the major parts of thehemp plant from both sexes. Figure 5 shows the 2D plots ofthe chromatographic separations of male bark (A), male leaf(C), female leaf (D), male hurd (E), female hurd (F) hempextract using chemiluminescence detection, and female leaf(B) using UV absorbance detection. The white points repre-sent peaks detected while the assigned letters correspond tothe respective cannabinoids as per Fig. 4.

At first sight, the general distribution of the cannabissample has a surprisingly high resemblance with that ofpreviously analyzed coffee samples by Mnatsakanyan and



Figure 5. 2D chemiluminesc-ence contour plots with CNfirst dimension and short C18

(50 mm) second dimension(A) Male Bark; (B) Female LeafUV; (C) Male Leaf; (D) FemaleLeaf; (E) Male Hurd; (F) FemaleHurd ESI1 Separation of femaleleaf extract on (a) Monolithicand (b) Short C18 column. Bothgradient increasing from 0 to100% methanol in 3 min andholding the organic phase forthe next 3 min. ESI2 Heart cutof (a) female leaf and (b) maleleaf extracts at 2.2 min from CNcolumn to short C18 column.

co-workers [25]. In their analysis, the separation obtainedfrom a CN first dimension and 150 mm C18 second dimen-sion were examined as three primary zones. The same ap-proach was performed here with the three zones listed asregion 1, region 2, and region 3 on Fig. 4 within the 2Dseparation plane. The two optimized dimensions showed dis-tinct retention behavior within specific regions of the sepa-ration space yielding separation of compounds, which were

co-eluting beforehand. For example, the compounds in re-gion 1 were poorly separated on the first dimension but havehad much greater retention and subsequent separation inthe second dimension. Region 2 corresponds to the broadband that was observed in the original separations. While lit-tle separation of those compounds was observed on the C18

dimension; they were selectively separated on the CN phase.It is important to note that the cannabinoids of interest were



present in the top right hand portion of this region (Fig. 4).It is not surprising to observe most of the cannabinoids elut-ing in the same region as they essentially have the sameC21 terpenophenolic structure with minor changes in theirfunctional groups. Finally, the compounds in region 3 werestrongly retained on both phases.

Although the overall distribution of the cannabis con-stituents in each sample is similar, there were subtle dif-ferences in their chemical composition. The female hurdhad the least number of components; lower signal inten-sity was also noted. As expected, the leaves were found tocontain the greatest number of components with 125 peaksdetected in the female leaf and 123 peaks in the male leafusing UV absorbance, 11 of which have been identified. An-other distinct feature of the leaves’ profile is the presence ofCBCV/CBDV/CBLV (peak b), while CBC/CBD/CBL (peak e)and CBN (peak d) were found in all the extracts.

A comparison between Fig. 5 (b) and (d) indicates sub-stantial differences in the UV absorbance and chemilumi-nescence detection of the female leave separations. A majordifference between UV and CL detection is the presence ofhigh intensity peaks in the bottom left corner of the UV plot.Although present in a large quantity in the cannabis sample,those peaks are not one of the 17 cannabinoids presented inFig. 3(A) and are not of specific interest to this study. How-ever it is important to note that this area of separation is com-monly of importance in 2D-HPLC separations and would beof significance in a chemical fingerprinting study of cannabissamples.

The use of chemiluminescence detection offers detectionselectivity toward the reducing agents present in the extractand allows a screening of the cannabinoids. We have previ-ously shown that the signal intensity generated by an analytewith acidic potassium permanganate chemiluminescence isrelated to the antioxidant activity of the molecules [19]. Theinitial driver for this proposal was to identify the key bioactiveconstituents in the hemp extract and their ability to act as an-tibacterial agents. Many antioxidants and cannabinoids havebeen shown to offer antibacterial activity and it is possiblethat the cannabinoids that have been selectively determinedwith the chemiluminescence reagent here may be directly dis-playing this bioactivity [30, 31]. The detection of fewer peakswith chemiluminescence may thus provide the potential de-termination of chemically active compounds in the sample.To fully realize this hypothesis however, a detailed study onthe structure–function relationship and chemiluminescencesignal intensity would need to be performed in conjunctionwith a cellular bioactivity test. This 2D LC technique is anoriginal approach to overcome frequently faced issues dur-ing cannabinoids analysis in both GC and uni-dimensionalHPLC [2].

4 Concluding remarks

This 2D-HPLC technique is an original approach to overcomefrequently faced issues during cannabinoids analysis in both

GC and uni-dimensional HPLC. The combination of 2D sep-aration with both UV absorbance and enhanced acidic potas-sium permanganate chemiluminescence detection is a bigleap forward in profiling technologies for cannabinoids, offer-ing comprehensive information about complex matrix char-acterization. This versatile separation technique can also beused to address current uncertainties about complex cannabi-noid chemistry for diverse purposes in a range of areas in-cluding taxonomical species identification, and in forensicclassification (fiber/drug type), identification, and individual-ization (source tracing). Further application of this procedureto phytochemical and pharmacological studies may bring theselectivity needed for the accurate identification of cannabi-noid profiles.

X.C., N.B., and P.S. would like to acknowledge the sup-port of Australian Research Council Linkage Project FundingLP130100681. J.P. thanks RMIT University and the Common-wealth Scientific and Industrial Research Organisation (CSIRO)for the award of a PhD scholarship (grant number 13/03634).

The authors have declared no conflict of interest.

5 References

[1] Hazekamp, A., Fischedick, J. T., Dı́e, M. L., Lubbe, A.,Ruhaak, R. L., Comprehensive Natural Products II 2010,3, 1033–1084.

[2] Raharjo, T. J., Verpoorte, R., Phytochem. Anal. 2004, 15,79–94.

[3] Elsohly, M. A., Slade, D., Life Sci. 2005, 78, 539–548.

[4] Mechoulam, R., Ben-Zvi, Z., J. Chem. Soc. D 1969, 7,343–344.

[5] Turner, C. E., ElSohly, M. A., Boeren, E. G., J. Nat. Prod.1980, 43, 169–234.

[6] ElSohly, M. A., Marijuana and the Cannabinoids, Hu-mana Press, New Jersey 2007.

[7] Hillig, K. W., Mahlberg, P. G., Am. J. Bot. 2004, 91, 966.

[8] Hazekamp, A., Proefschrift Universiteit Leiden 2007.

[9] Callaway, J. C., Euphytica 2004, 140, 65–72.

[10] Dussy, F. E., Hamberg, C., Luginbuhl, M., Schwerzmann,T., Briellmann, T. A., Forensic Sci. Int. 2005, 149, 3–10.

[11] Hazekamp, A., Peltenburg, A., Verpoorte, R., Giroud, C.,J. Liq. Chromatogr. Relat. Technol. 2005, 28, 2361–2382.

[12] De Backer, B., Debrus, B., Lebrun, P., Theunis, L., Dubois,N., Decock, L., Verstraete, A., Hubert, P., Charlier, C.,J. Chromatogr. B: Anal. Technol. Biomed. Life Sci. 2009,877, 4115–4124.

[13] Rustichelli, C., Ferioli, V., Vezzalini, F., Rossi, M. C., Gam-berini, G., Chromatographia 1996, 43, 129–134.

[14] Stolker, A. A. M., Schoonhoven, J. V., Vries, A. J. D.,Bobeldijk-Pastorova, I., Vaes, W. H. J., Berg, R. V. D.,J Chromatogr A 2004, 1058, 143–151.

[15] Guiochon, G., J Chromatogr A 2006, 1126, 6–49.

[16] Cohen, S. A., Schure, M. R., Multidimensional LiquidChromatography John Wiley & Sons, Hoboken 2008.



[17] Bassenasse, D.N., Conlan, X. A., Barnett, N. W., Steven-son, P. G., J. Sep. Sci. 2015, DOI:10.1002/jssc.201500054

[18] Stevenson, P. G., Bassenasse, D.N., Barnett, N. W., Con-lan, X. A., J. Sep. Sci. 2013, 36, 3503–3510

[19] Conlan, X. A., Stupka, N., McDermott, G. P., Bar-nett, N. W., Francis, P. S., Anal. Methods 2010, 2,171–173.

[20] Terry, J. M., Adcock, J. L., Olson, D. C., Wolcott, D. K.,Schwanger, C., Hill, L. A., Barnett, N. W., Francis, P. S.,Anal. Chem. 2008, 80, 9817–9821.

[21] Fairchild, J. N., Horvath, K., Gooding, J. R., Campagna,S. R., Guiochon, G., J. Chromatogr. A 2010, 1217, 8161–8166.

[22] Cohen, S. A., Schure M. R., Multidimensional LiquidChromatography, Wiley & Sons, Hoboken 2008.

[23] Shalliker, R. A., Kavanagh, P. E., Russell, I. M., J. Chro-matogr. A 1991, 558, 440–445.

[24] Jandera, P., Guiochon, G., J. Chromatogr. A 1991, 588,1–14.

[25] Mnatsakanyan, M., Stevenson, P. G., Conlan, X. A., Fran-cis, P. S., Goodie, T. A., McDermott, G. P., Barnett, N. W.,Shalliker, R. A., Talanta 2010, 82, 1358–1363.

[26] Gray, M. J., Dennis, G. R., Slonecker, P. J., Shalliker, R.A., J. Chromatogr. A 2004, 1041, 101–110.

[27] Liu, Z. D. G. P., H., Anal. Chem. 1995, 67, 3840–3845.

[28] Stevenson, P. G. G., G., J. Chromatogr. A 2012, 1247,57–62.

[29] Holland, B. J., Deakin University, Waurn Ponds 2011, p.58.

[30] Appendino, G., Gibbons, S., Giana, A., Pagani, A., Grassi,G., Stavri, M., Smith, E., Rahman, M. M., J. Nat. Prod.2008, 71, 1427–1430.

[31] Caturla, N., Vera-Samper, E., Villalain, J., Mateo, C. R.,Micol, V., Free Radical Biol. Med. 2003, 34, 648–662.


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