Lobed Nuclei still count as One nucleus?

Lobed Nuclei still count as One nucleus?

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Do the Lobed Nuclei of immune cells (such as Megakaryocytes) still count as one nucleus?

Yes. Lobation is when a nucleus deforms, but it is still a single compartment. How the nucleus deforms can be helpful in roughly determining the cell type by visual inspection.


Monocytes are a type of leukocyte, or white blood cell. They are the largest type of leukocyte and can differentiate into macrophages and myeloid lineage dendritic cells. As a part of the vertebrate innate immune system monocytes also influence the process of adaptive immunity. There are at least three subclasses of monocytes in human blood based on their phenotypic receptors.


There are four types of granulocytes (full name polymorphonuclear granulocytes): [3]

Except for the mast cells, their names are derived from their staining characteristics for example, the most abundant granulocyte is the neutrophil granulocyte, which has neutrally staining cytoplasmic granules.

Neutrophils Edit

Neutrophils are normally found in the bloodstream and are the most abundant type of phagocyte, constituting 60% to 65% of the total circulating white blood cells, [4] and consisting of two subpopulations: neutrophil-killers and neutrophil-cagers. One litre of human blood contains about five billion (5x10 9 ) neutrophils, [5] which are about 12–15 micrometres in diameter. [6] Once neutrophils have received the appropriate signals, it takes them about thirty minutes to leave the blood and reach the site of an infection. [7] Neutrophils do not return to the blood they turn into pus cells and die. [7] Mature neutrophils are smaller than monocytes, and have a segmented nucleus with several sections(two to five segments) each section is connected by chromatin filaments. Neutrophils do not normally exit the bone marrow until maturity, but during an infection neutrophil precursors called myelocytes and promyelocytes are released. [8]

Neutrophils have three strategies for directly attacking micro-organisms: phagocytosis (ingestion), release of soluble anti-microbials (including granule proteins), and generation of neutrophil extracellular traps (NETs). [9] Neutrophils are professional phagocytes: [10] they are ferocious eaters and rapidly engulf invaders coated with antibodies and complement, as well as damaged cells or cellular debris. The intracellular granules of the human neutrophil have long been recognized for their protein-destroying and bactericidal properties. [11] Neutrophils can secrete products that stimulate monocytes and macrophages these secretions increase phagocytosis and the formation of reactive oxygen compounds involved in intracellular killing. [12]

Neutrophils have two types of granules primary (azurophilic) granules (found in young cells) and secondary (specific) granules (which are found in more mature cells). Primary granules contain cationic proteins and defensins that are used to kill bacteria, proteolytic enzymes and cathepsin G to break down (bacterial) proteins, lysozyme to break down bacterial cell walls, and myeloperoxidase (used to generate toxic bacteria-killing substances). [13] In addition, secretions from the primary granules of neutrophils stimulate the phagocytosis of IgG antibody-coated bacteria. [14] The secondary granules contain compounds that are involved in the formation of toxic oxygen compounds, lysozyme, and lactoferrin (used to take essential iron from bacteria). [13] Neutrophil extracellular traps (NETs) comprise a web of fibers composed of chromatin and serine proteases that trap and kill microbes extracellularly. Trapping of bacteria is a particularly important role for NETs in sepsis, where NET are formed within blood vessels. [15]

Eosinophils Edit

Eosinophils also have kidney-shaped lobed nuclei (two to four lobes). The number of granules in an eosinophil can vary because they have a tendency to degranulate while in the blood stream. [16] Eosinophils play a crucial part in the killing of parasites (e.g., enteric nematodes) because their granules contain a unique, toxic basic protein and cationic protein (e.g., cathepsin [13] ) [17] receptors that bind to IgE are used to help with this task. [18] These cells also have a limited ability to participate in phagocytosis, [19] they are professional antigen-presenting cells, they regulate other immune cell functions (e.g., CD4+ T cell, dendritic cell, B cell, mast cell, neutrophil, and basophil functions), [20] they are involved in the destruction of tumor cells, [16] and they promote the repair of damaged tissue. [21] A polypeptide called interleukin-5 interacts with eosinophils and causes them to grow and differentiate this polypeptide is produced by basophils and by T-helper 2 cells (TH2). [17]

Basophils Edit

Basophils are one of the least abundant cells in bone marrow and blood (occurring at less than two percent of all cells). Like neutrophils and eosinophils, they have lobed nuclei however, they have only two lobes, and the chromatin filaments that connect them are not very visible. Basophils have receptors that can bind to IgE, IgG, complement, and histamine. The cytoplasm of basophils contains a varied amount of granules these granules are usually numerous enough to partially conceal the nucleus. Granule contents of basophils are abundant with histamine, heparin, chondroitin sulfate, peroxidase, platelet-activating factor, and other substances.

When an infection occurs, mature basophils will be released from the bone marrow and travel to the site of infection. [22] When basophils are injured, they will release histamine, which contributes to the inflammatory response that helps fight invading organisms. Histamine causes dilation and increased permeability of capillaries close to the basophil. Injured basophils and other leukocytes will release another substance called prostaglandins that contributes to an increased blood flow to the site of infection. Both of these mechanisms allow blood-clotting elements to be delivered to the infected area (this begins the recovery process and blocks the travel of microbes to other parts of the body). Increased permeability of the inflamed tissue also allows for more phagocyte migration to the site of infection so that they can consume microbes. [19]

Mast cells Edit

Mast cells are a type of granulocyte that are present in tissues [3] they mediate host defense against pathogens (e.g., parasites) and allergic reactions, particularly anaphylaxis. [3] Mast cells are also involved in mediating inflammation and autoimmunity as well as mediating and regulating neuroimmune system responses. [3] [23] [24]

Granulocytes are derived from stem cells residing in the bone marrow. The differentiation of these stem cells from pluripotent hematopoietic stem cell into granulocytes is termed granulopoiesis. Multiple intermediate cell types exist in this differentiation process, including myeloblasts and promyelocytes.

Granule contents Edit

Examples of toxic materials produced or released by degranulation by granulocytes on the ingestion of microorganisms are:

Granulocytopenia is an abnormally low concentration of granulocytes in the blood. This condition reduces the body's resistance to many infections. Closely related terms include agranulocytosis (etymologically, "no granulocytes at all" clinically, granulocyte levels less than 5% of normal) and neutropenia (deficiency of neutrophil granulocytes). Granulocytes live only one to two days in circulation (four days in spleen or other tissue), so transfusion of granulocytes as a therapeutic strategy would confer a very short-lasting benefit. In addition, there are many complications associated with such a procedure.

There is usually a granulocyte chemotactic defect in individuals suffering from type 1 diabetes mellitus.

Research suggests giving granulocyte transfusions to prevent infections decreased the number of people who had a bacterial or fungal infection in the blood. [25] Further research suggests participants receiving therapeutic granulocyte transfusions show no difference in clinical reversal of concurrent infection. [26]

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Results and discussion

First, we chose to use the Arabidopsis root to validate the effectiveness of our protoplasting-free single-nucleus RNA sequencing approach because of the well-studied cell types [27] and the rich resource of single-cell data [11,12,13,14,15,16] of this tissue. We directly isolated nuclei by sorting from homogenized root tips of 10-day-old Arabidopsis seedlings without protoplasting (Additional file 1: Fig. S2). The nuclei were fed to the 10x Genomics Chromium platform to obtain full-length cDNA templates labeled with nucleus-specific barcodes, which are subsequently divided into two equal parts and used for constructing Illumina short-read and Nanopore long-read libraries, respectively (Fig. 1a).

From the Illumina library, we obtained a total of 1186 single-nucleus transcriptomes covering 18,913 genes, with median genes/nucleus at 810 and median UMIs/nucleus at 1131. It is worth noting that the proportion of intron-containing mRNAs is extremely high in plant nucleus—54% compared to less than 2% in total RNAs [28] (Fig. 1b). After generating the cell-gene abundance matrix from Illumina data, we utilized an unbiased graph-based clustering method Louvain [29] and identified 14 distinct cell clusters (Fig. 1c). We then applied a set of cell type-specific marker genes provided in a recent massive single-cell study of Arabidopsis roots [17] to annotate each cluster (see the “Methods” section, Additional file 2: Table S1). We were able to assign cell types to all 14 clusters and identified 10 major root cell types previously reported (Fig. 1c, Additional file 1: Fig. S3), with the signature transcripts for each cell type enriched in the corresponding cluster (Fig. 1d, Additional file 1: Fig. S4). Consistent with previous reports [11,12,13,14,15,16], we also noticed that some cell types from our result are composed of multiple clusters, such as stem cell niche (clusters 1, 4, and 12), mature non-hair (clusters 2 and 6), and endodermis (clusters 5 and 8) (Fig. 1c), demonstrating additional heterogeneity (subcell types) within cell types. Moreover, we found the exact same subcell type marker genes of endodermis are enriched in each of its corresponding subcell types as shown in Zhang et al. [15] (Additional file 1: Fig. S5), demonstrating the robustness of our single-nucleus data. In addition, we used the Scanorama algorithm [30] to compare our dataset with several recently published root single-cell datasets from protoplasts [11, 12, 14,15,16]. The expression abundance matrix from our single-nucleus dataset closely resembles the protoplasting-based single-cell dataset generated from the same tissue (10-day seedling, 0.5 mm primary root tips) [15] (Fig. 2a, b). Taken together, we demonstrated that transcriptomes of the single nucleus are sufficient for cell type identification and can be used as a reliable alternative to protoplasts.

Dataset generated by flsnRNA-seq is consistent with protoplast-based scRNA-seq. a Heatmap represents alignment score between the single-nucleus data and single-cell datasets generated from 10x Genomics platform. Alignment score is calculated by Scanorama [30]. Higher alignment score indicates higher similarity between a pair of datasets. b Pairwise integration of two single cell/nucleus datasets. The batch effect is removed by Scanorama. The expression matrix is downsampled to the same dimension as the single-nucleus data

As to the Nanopore data analysis, a key challenge is that the relatively low sequencing accuracy of Nanopore (

95% per base) makes it difficult to correctly recognize the cell barcodes and UMI information on each Nanopore read. To solve this problem, Lebrigand et al. developed a method named Sicelore to use Illumina short reads generated from the same cDNA library as the guide to allocate Nanopore reads [24]. Sicelore searches for both polyA and adapter sequence and defines the region between these two as the potential barcode and UMI. However, this algorithm relies on the recognition of polyA tail sequence generated by the Nanopore basecalling software, which tends to severely underestimate the length of polyA tail [31]. We tried to further improve Sicelore by developing a polyA-independent algorithm (named snuupy), which searches for cell barcodes and UMIs in the unmapped region of Nanopore reads (see the “Methods” section and Additional file 1: Fig. S1, Fig. 3a). As the result, snuupy recovers 20% more reads from our Nanopore data compared to using Sicelore [24] (Fig. 3b). After snuupy processing, we obtained 1169 long-read single-nucleus transcriptomes from Nanopore data (compared to the 1186 from Illumina data). The median UMI counts per nucleus (729) and the median gene counts per nucleus (563) from Nanopore data are

70% of the Illumina count, respectively, and highly consistent in all nuclei (Fig. 3c). The clustering result using Nanopore abundance matrix closely resembles the one generated by Illumina data (Fig. 3d, e), suggesting that Nanopore data itself is sufficient for cell type classification, consistent with a recent large-scale single-cell analysis in human and mouse cells performed entirely with Nanopore data [24, 25].

Snuupy assigns cell barcodes and UMIs for Nanopore reads according to the information from Illumina data. a Flowchart shows the difference between snuupy and Sicelore. b Overlap between snuupy and Sicelore allocated reads. c Numbers of UMIs (left) and genes (right) detected in each nucleus from the Illumina and Nanopore data. d UMAP visualization of the root cell types clustered using abundance information from the Nanopore single-nucleus data. The cell color is the same as in Fig. 1c. e UMAP visualization of the integration of two datasets. The batch effect is removed by Scanorama [30]. Alignment score is calculated by Scanorama and in the range from 0 to 1. Higher alignment score indicates a higher similarity between a pair of datasets

The single-nucleus long-read Nanopore library provides isoform-level information such as splicing and APA, compared to Illumina library which only captures abundance information of transcripts. Therefore, we generated two additional isoform matrices to track splicing and APA in single nucleus, respectively (Fig. 4a and Additional file 1: Fig. S6), and combined them with the Illumina abundance matrix for a multilayer clustering, to test if these extra layers of information could improve cell type classification. Indeed, we found that the original cluster 2 (mature non-hair) and cluster 10 (cortex) from Illumina data (Fig. 1c) can be further separated into two subcell type clusters after the multilayer clustering (Fig. 4a). As an example, from the Illumina data, transcripts of AT3G19010 are present in both subcell type 2.1 and 2.2 (Fig. 4b, c), while the Nanopore data revealed a large difference at the splicing level of this gene between the two sub-clusters, with the second intron largely unspliced in subcell type 2.2 (Fig. 4d). It is worth noting that, JAZ7, the top 1 enriched gene in cluster 2.2 (Fig. 4e), can regulate splicing during jasmonate response [32], implying a fascinating potential of cell type-specific regulation of splicing that could be investigated with flnsRNA-seq.

Nanopore long-read single-nucleus RNA-seq improves cell type identification. a Multilayer matrices combining Illumina abundance matrix with Nanopore splicing and APA information improve cell type identification. b, c Genome-browser plot of Illumina reads (b) and Nanopore reads (c) aligned to gene AT3G19010. The second intron of AT3G1910 shows different splicing patterns between cluster 2.1 and cluster 2.2. The red arrowhead indicates the second intron. Differences in splicing patterns between two clusters were tested using Fisher exact test, and the corresponding p value is lower than 0.001. The red bar at the 3′ end of Nanopore reads (blue) indicates the Poly(A) tail. d UMAP visualization shows the abundance distribution of AT3G19010 as well as the differential splicing of the second intron between cluster 2.1 and cluster 2.2. e The top 25 genes enriched in cluster 2.2 are ranked by enriched score compared to cluster 2.1 (upper panel) and UMAP visualization shows the abundance distribution of the most enriched gene JAZ7 (lower panel). The enriched score is calculated using rank_genes_groups function of Scanpy. The red arrowhead indicates the most enriched gene in cluster 2.2

After establishing flsnRNA-seq using well-documented root tissue, we applied this method to investigate other tissues that have not been previously characterized at the single-cell level due to difficulties in obtaining corresponding protoplasts. In flowering plants, seed development is initiated by double fertilization, during which egg cell and central cell fuse with sperm cells respectively to form embryo and endosperm [33]. The endosperm is embedded in the seed coat and responsible for providing nutrients from maternal parent to developing embryo [34, 35]. In Arabidopsis endosperm, the primary nucleus formed after fertilization undergoes several rounds of rapid nuclear divisions without cytokinesis, resulting in a multinucleate cell termed syncytium, which later cellularized and differentiated into three endosperm domains: the micropylar, central peripheral, and chalazal [36, 37] (Fig. 5a). Cellularization of the syncytium is critical for embryo viability [34], and this process is initiated when the embryo reaches the heart stage, starting from the micropylar domain and gradually proceeds to the central periphery in a wavelike pattern and eventually reach the chalazal zone [38]. Transcriptomes from various developing stages of bulked endosperm have been well-documented using microarray or RNA-seq [39,40,41,42,43,44,45] however, endosperm has yet to be characterized at the single-cell level due to technical challenges in generating protoplasts from the endosperm.

flsnRNA-seq captures the variation in intron retention levels of different clusters. a UMAP visualization of clustering result using Illumina single-nucleus data (left panel), and cartoon illustration of major cell types in Arabidopsis endosperm at heart stage (right panel). b UMAP visualization of incompletely spliced ratio calculated by Nanopore full-length reads. c Barplot visualization of the incompletely spliced ratio of each cluster. Differences in incompletely spliced ratios between each cluster to all other clusters were tested using a one-sided Kolmogorov-Smirnov test. “***” denotes that the p value is lower than 0.001. d Quantification of nuclei with each cell type per cluster. The number represents nucleus counts and the color represents the proportion of cell types in each cluster. e GO term enrichment analysis of all 93 enriched genes for cluster 4. Only cellular component terms are plotted. “*” and “***” denote that the adjusted p value is lower than 0.05 and 0.001, respectively

Here, we applied flsnRNA-seq to the multinucleate endosperm isolated from the heart-stage ovules of Arabidopsis and generated both the Illumina and Nanopore 10x libraries. We obtained 576 nuclei from Illumina data, with the median genes/nucleus at 645 and the median UMIs/nucleus at 853. All 576 nuclei were captured by the Nanopore library, with the median genes/nucleus at 300 and the median UMIs/nucleus at 362 (Additional file 1: Fig. S7). Based on the Illumina abundance matrix, we identified six clusters using the Louvain method (Fig. 5a). Next, we used Nanopore full-length transcript data to analyze retained introns in each nucleus and found that the nuclei from cluster 4, a major cluster that accounts for 14% of the total nuclei, exhibits a distinct high ratio of incompletely spliced transcripts (Fig. 5b, c). Several previous studies have established that increased accumulation of intronic reads is an indicator of transcription activation [46, 47], and the ratio of unspliced precursor and spliced mature transcripts have been widely used to estimate RNA velocity [48, 49]. The high ratio of incompletely spliced transcripts in this particular cluster of endosperm nuclei may reflect delayed pre-mRNA decay, disrupted intron turnover rate, or a global activation of transcription. Next, we assigned the cell type of each nucleus using previously reported cell type-enriched genes at the heart stage [50, 51] and found that the majority of nuclei in cluster 4 are annotated as micropylar endosperm (Fig. 5d). In addition, gene ontology (GO) analysis of the genes upregulated in cluster 4 found all enriched cellular component terms are membrane related (Fig. 5e, Additional file 2: Table S3), suggesting these nuclei are poised to forming the cellular membrane and entering the cellularization stage, consistent with the previous observation that cellularization of the Arabidopsis endosperm is first initiated at the micropylar endosperm [38, 52]. Hence, our method identified a unique cluster of endosperm nuclei with a high proportion of incompletely spliced transcripts, and further investigation could determine whether this is due to the increase in transcription or delay in splicing.

Lobed or segmented nuclei

Granulocytes of the immune system

Vertebrate immune systems contain a variety of white cells from the myeloid lineage, termed granulocytes for their cytoplasmic appearance under haematoxylin and eosin dye. The granulocytes have been commonly recognised and distinguished histologically by their nuclear shapes and sizes. They contain multi-lobed nuclei, each lobe connected by a short region of nucleoplasm (Fig.  1b ). Of the granulocytes, eosinophils have the fewest lobes. Their bi-lobed nucleus together with their intense eosin staining means they are often described to histology students as a sunburned face wearing dark sunglasses. Much of the variation within each cell type is found in the number of lobes an increased lobe number is termed hypersegmentation. Hypersegmentation of eosinophils is rare, but has been seen in acute eosinophilic pneumonia, with lobe number increased to three or four lobes (Maeno et al. 2000), and could be linked to stimulation with lymphokines (Chihara and Nakajima 1989). In basophils, hypersegmentation is also rare but has been occasionally observed (Xu 2014). However, most of the studies of granulocyte nuclear structure have been performed on the neutrophils.


Mammalian neutrophils𠅊nd avian or reptilian heterophils (Claver and Quaglia 2009)—have segmented, multi-lobed nuclei, usually containing between two and five lobes, separated by thin filaments of nucleoplasm with little to no internal chromatin. The lobed structure develops from a spherical myelocyte precursor, gradually increasing the number and prominence of lobes through the concave metamyelocyte and band cell stages to the mature neutrophil (Fig.  1b ).

Chromosome painting and 3D analysis have shown that most chromosomes are randomly distributed within neutrophil lobes, but the organisation can change upon bacterial stimulation (Yerle-Bouissou et al. 2009 Mompart et al. 2013). Within each lobe, the chromatin organisation follows a general gene-density based arrangement, in which the gene-poor chromatin is located towards the nuclear periphery, and gene-dense chromatin more internal (H࿋ner et al. 2015). Curiously, the inactive X chromosome in women is frequently found in a terminal lobe, often with a distinct 𠆍rumstick’ appearance (Karni et al. 2001), and it appears that the position of the inactive X within the precursor myelocyte may determine the polarity of the neutrophil. It remains unknown how polarity is determined in XY neutrophils.

Hypersegmentation of neutrophils, to six or more lobes, is associated with megaloblastic anaemias, such as result from deficiencies in Vitamin B12 and folic acid, and iron deficiency anaemia (Westerman et al. 1999). It is also associated with Boucher-Neuhäuser syndrome (Umehara et al. 2010 Koh et al. 2015), a lipid metabolic defect. In rats, vitamin A deficiency caused hypersegmentation, linked to a requirement of retinoids for differentiation of promyelocytes to mature neutrophils (Twining et al. 1996). Consequently, there are clearly many pathways that contribute to the establishment of a lobed nuclear morphology. What though is it for?

Functional significance of a lobed nucleus

It is thought that the lobular arrangement makes the nucleus easier to deform and, hence, help the neutrophils pass through small gaps in the endothelium and extracellular matrix more easily (Hoffmann et al. 2007) granulocytes with defects in lamin B receptors (a component of the inner nuclear membrane) are unable to adopt a normal segmented shape, have fewer lobes (Hoffmann et al. 2002), and are poorer at passing through these small spaces. Neutrophils also have a higher variability in the length of the linker DNA between nucleosomes than T-lymphocyte populations (Valouev et al. 2011), pointing to increased chromatin flexibility.

However, neutrophils are not the only migratory cell in circulation circulating monocytes, for example, have a lobed nucleus but, as described below, the lobes are larger and fewer. Monocytes are also flexible enough to enter tissues, whereupon they differentiate into various other cell types including macrophages. Indeed, comparisons of the migration of monocytes and neutrophils suggest that the monocytes are at least equally flexible when penetrating basement membranes, and that neutrophil migration is aided by reorganisation of the extracellular matrix via proteolytic cleavage of laminins (Voisin et al. 2009). The circulating fibrocytes and lymphocytes mentioned below are also migratory and have spindle-shaped and spherical nuclei, respectively.

Consequently, while the lobular shape of neutrophils may aid migration, is not strictly necessary for migration. Why then should neutrophils adopt lobes, when other cells do not? Perhaps the answer lies in the lifespan of the cells. The half life of a neutrophil in circulation is about 6 h (Summers et al. 2010). Though circulating monocytes live only a couple of days, macrophages may live for months in a tissue, as can lymphocytes.

The granulocytes have lower lamin protein content than macrophages or monocytes—predominantly a loss of the lamins A and C, with an increase in lamin B (Hoffmann et al. 2007). The lamin proteins, as described in more detail later, provide structural support to the nucleus, and protect against damage from mechanical stresses. Particularly, the ratio of lamin A:B balances the stiffness of the nucleus against its elasticity (Shin et al. 2013). Correspondingly, defects in the lamins associated with normal aging affects nuclear shape in all the granulocytes (Scaffidi et al. 2005 Chan et al. 2010), a result of changes to the stiffness and structure of the nuclear lamina. These age-related structural defects are also seen in laminopathies such as Hutchinson-Gilford Progeria Syndrome (Worman and Courvalin 2005).

Furthermore, rats treated with cyclophosphamide (a DNA cross-linker that disrupts DNA replication) have hypersegmented tetraploid neutrophils in their blood (Kotelnikov et al. 1988). The underlying mechanism driving hypersegmentation seems to be both failures during DNA synthesis and DNA damage or loss of nuclear structural integrity. Consequently, it appears that the extra flexibility of neutrophil nuclei comes at the cost of lowering their lifespan (Harada et al. 2014), a cost that other, longer-lived cell types cannot bear.

Neutrophil extracellular traps

Neutrophils are capable of a form of cell death termed ‘NETosis’. They produce meshes of chromatin complexed with cytoplasmic proteins, termed Neutrophil Extracellular Traps (NETs), which capture bacteria (Brinkmann and Zychlinsky 2007 Brinkmann and Zychlinsky 2012). Such traps have been seen in orthologous cell types across vertebrates. During the process of NET formation, the nucleus loses its lobular structure, and the chromatin decondenses. The nuclear and cell membranes break down, releasing the NET into the extracellular space over 𢏁𠄴 h. In particular circumstances, such as in response to Staphylococcus aureus, neutrophils may be able to generate NETs without lysis of the cell, by generating chromatin-filled vesicles that rupture after budding, a process that can happen in only minutes to an hour (Pilsczek et al. 2010).

Interestingly, NETs (or equivalents) can be produced by other leukocytes in addition to neutrophils (Goldmann and Medina 2013), such as mast cells and eosinophils. It remains unclear whether the lobular structure of the granulocyte nucleus is relevant for the formation of NETs (Veda 2011), and studies are needed to test the effects of the reduced structural stability of the nucleus on the speed or ease with which NETs can be formed.

Monocytes and macrophages

Monocytes have a bilobed nucleus (Fig.  1c ), which frequently presents in tissue sections and blood smears as a U- or kidney-shaped nucleus. The lobed structure arises in promonocytes, where an initial spherical nucleus acquires an indentation that develops into the separation of the lobes (Fawcett 1970). The reason for the lobed structure is still unclear perhaps it helps with the flexibility of the nucleus, but leaves the nucleus less susceptible to damage than the highly segmented granulocytes.

The nucleus generally becomes more rounded following recruitment into tissues and further differentiation into a variety of macrophages and other cell types (Mosser and Edwards 2008). At high resolution, a clear difference is observable in the chromatin distribution within the nuclei. Chromatin domains within monocytes are aggregated into clusters, with channels and spaces between them. In monocytes𠅊nd indeed granulocytes—the channels and spaces within the nucleus are large, and may facilitate chromatin deformation upon migration (H࿋ner et al. 2015).

Even after differentiation into a macrophage, the cell nucleus can undergo extensive deformation in response to environmental conditions. Examples of nuclear reshaping of macrophages can be seen in electron microscopy images (Sato-Nishiwaki et al. 2013), and the nucleus is both displaced with the cell and reshaped from round to kidney-shaped in response to Bacillus anthricis edema toxin (Trescos et al. 2015). It is worth noting that macrophages remain functionally plastic—they can change between roles with relative ease (Mosser and Edwards 2008), and perhaps the readily deformable nucleus facilitates this via impacts on transcriptional regulation.

Many questions remain about these cells. Individual macrophages can fuse into giant macrophages (see Fig.  1c ), thought to improve the efficiency of phagocytosis (McNally and Anderson 2011). Electron microscopy images show dense packing and distortion of abutting nuclei in giant cells (Sutton and Weiss 1966), but how do these shapes affect function and what is the relevance of nuclear position within these cells, such as the Langhans-type giant cells in which nuclei form a horseshoe around the periphery?


Megakaryocytes are the precursor cells from which platelets will develop by fragmentation of the cytoplasm. Their large multilobed nuclei are produced by successive rounds of endomitosis—that is, cell division in which the mitotic cycle stops during anaphase, skipping telophase and cytokinesis (Patel et al. 2005). This results in a large nucleus with a variable DNA content from 4 to 128 N.

In contrast to granulocytes, the lobes appear clustered, like a bunch of grapes, rather than separated by strands. Furthermore, there appears to be a difference in chromosomal segregation patterns between high and low ploidy cells (Papadantonakis et al. 2008). Although the nuclei are variable in morphology between cells, there are some clear morphological appearances that can be used to identify pathologies. For example, chronic myeloproliferative disorders are often accompanied by irregularities in morphology, and increased variation in lobe number (Ballarò et al. 2008), probably a symptom of disruptions to the structure of the nuclear envelope. Multinucleated megakaryocytes, as can arise in dysplasias, appear to arise from a further progression through the mitotic cycle (Münch et al. 2011).

It remains uncertain what the functional relevance of the ploidy or lobulation is in megakaryocytes they exhibit functional gene expression amplification resulting from the polyploidy, but studies attempting to link platelet formation with ploidy and morphology have yielded inconclusive results to date (Machlus and Italiano 2013). Another common mammalian polyploid nucleus, that of the hepatocyte, is not lobed, but tends only to reach 8 N. Consequently, it remains unclear whether the lobulation is a physical response to the greater ploidy, or a result of inherited differentiation or programming pathways shared with the granulocyte lineages.

Essay on the Life Cycle of Ginkgo Biloba | Gymnosperms | Botany

Read this essay to learn about the life cycle of ginkgo biloba, explained with the help of suitable diagrams.

Sporophyte of Ginkgo Biloba:

Ginkgo biloba is a tall deciduous tree (up to 30 m height) giving rise to a very irregular pattern of branching. The branching is restricted to the upper part of the stem, thus giving the tree an ex-current pattern of growth. The branches are dimorphic, bearing two types of shoot long shoots and dwarf shoots (Fig. 1.35).

The leaves also show dimorphism, those present on the long shoot are deeply lobed, while those on the dwarf shoots are either not so deeply lobed or may be entire. G. biloba has a tap root system which possesses strong lateral roots that penetrate deep into the soil.

The plants are dioecious. The male and female trees are only distinguishable after the production of reproductive structures on them.


Ginkgo biloba reproduces sexually. Ginkgo is dioecious. Morphologically, the male and the female plants are indistinguishable before the formation of reproductive organs. However, the male and the female trees can be distinguished cytologically at an early stage.

In the female plant, out of the 24 chromosomes, four are satellite-bearing chromosomes, while in male plant three chromosomes are with satellite. Like cycads, the reproductive structures of Ginkgo are most primitive among living seed plants.

Male strobili develop in the axil of leaves or on the sides of petiole bases present on dwarf shoots (Fig. 1.40A). The male strobilus is a loose structure and consists of a stalked central axis (2-3 cm in length) on which many microsporo­phylls are arranged spirally. A male strobilus looks like a catkin inflorescence of angiosperms (Fig. 1.40 A).

Each microsporophyll has a long slender stalk terminating into a knob-like portion or hump. It bears two pendent microsporangia (Fig. 1.40B). The hump is always provided with a mucilage duct.

The morphological nature of the hump is still an open question. According to some botanists, the hump represents an abortive sporangium and the mucilage cavity represents abortive sporogenous tissue. In rare instances, a microsporophyll bears three sporangia.

The microsporangia develops from a hypo- dermal archesporial cell of the microsporophyll. A mature microsporangium consists of a multi- layered wall, tapetum and microspore mother cells.

It is important to note that 1-3 hypodermal layers develop fibrous thickening representing an endothecium as in angiosperm. In this point, Ginkgo differs from other gymnosperms where the outermost layer of the microsporangium develops fibrous thickening representing an exothecium.

Inside the sporangium, each microspore mother cell produces four microspores or pollen grains by meiotic division. The pollen grain is oval-shaped with monosulcate aperture. Each grain is bounded by two concentric wall layers: the outer thick exine and the inner thin intine.

The exine is not continuous, being absent at the top apertural region. The dehiscence of sporan­gia takes place by longitudinal slits. At the time of dehiscence the sporangia are pulled apart due to the shrinkage of mucilage cavity of the hump.

Female Strobilus:

The female strobili develop in the axil of a leaf or a scale leaf present on dwarf shoots (Fig. 1.41). The female strobili are very much reduced structures. Each strobilus consists of a long stalk or peduncle that bifurcates at its apex and each branch usually bears a single sessile ovule (Fig. 1.42A).

Out of the two ovules, one remains viable while the other aborts at an early stage. Sometimes, both the ovules become viable and develop into seeds. The base of each ovule is surrounded by a fleshy cup called collar.

Most of the scientists believed that the female strobilus of G. biloba is a very reduced structure and the collar represents a reduced megasporo­phyll. Sometimes, the collar may grow into a leafy shoot — it supports this hypothesis.

Moreover, the peduncle bearing two ovules has four vascular traces, two in each ovule (Fig. 1.42B), thus representing the axis of a female stro­bilus bearing two megasporophylls. Some abnor­malities have also been observed where a single peduncle bears many ovules. In such conditions, the number of vascular traces in the peduncle would be twice the number of ovules.

Pankow and Sothman (1967) disagreed over the sporophyll nature of the collar. They consi­dered the ovules of Ginkgo biloba as cauline and terminal on lateral axis. They argued that the col­lar is devoid of any vascular supply and develops after the formation of integument.

The ovule of Ginkgo is similar to that of Cycas. The ovules are orthotropous, unitegmic and crassinucellate (with massive nuceller tissue). In the young ovule, the integument is free from the nucellus, except at the chalazal end, while in a matured ovule the integument is fused with the nucellus except at the apex due to the enlargement of the chalazal end (Fig. 1.42B).

The ovule has a massive beak-shaped nucellus (Fig. 1.42A). In a pre-pollinated ovule, the apical nucellar cells degenerates to form a deep pollen chamber and the pollination drop.

The single integument is differentiated into three layers:

The outer fleshy with numerous mucilage cavities, the middle stony and the inner fleshy, eventually becomes papery. The ovule is supplied with two vascular traces which enter into the inner fleshy layer reaching up to the free part of the nucellus.


A deeply situated cell of the nucellus is differentiated into a large megaspore mother cell.

The megaspore mother cell is surrounded by a well-developed spongy tissue. At a later stage, the spongy tissue cells adjacent to the enlarging gametophyte lose their cell walls. The mega­spore mother cell wall becomes thick due to the development of a double layered wall. Now, the megaspore mother cell undergoes meiotic divi­sion to form a liner tetrad of four megaspores.

The upper three megaspores degenerate, while the lowermost one becomes functional.

Gametophyte of Ginkgo Biloba:

The spores (either microspores or megas­pores) represent the first phase of gametophyte generation. The microspore or pollen grain is the male gametophyte, while the megaspore is the first stage of female gametophyte which deve­lops into a female gametophyte.

Development of Male Gametophyte before Polli­nation:

The pollen nucleus undergoes mitotic divi­sion to produce a small lens-shaped first prothalial cell towards the proximal end (opposite to the apertural side) and a large central cell on the distal end (Fig. 1.43A). The prothallial cell does not divide, whereas the central cell divides to form a second prothallial cell and an antheridial initial (Fig. 1.43B).

The first prothallial cell is ephemeral, while the second prothallial cell is persistent. The antheridial initial cuts off a small antheridial cell and a large tube cell (Fig. 1.43C). The pollen grains are released from the microsporangium at this 4-celled stage (2 pro­thallial cells, an antheridial cell and a tube cell).

Development of Male Gametophyte after Polli­nation:

The further development of male gameto­phyte starts within a month after pollination. The tube cell of the pollen comes out through the aperture in the form of a pollen tube. The tube proceeds towards the archegonium, penetrating the nucellar tissue of the ovule. The antheridial cell within the pollen tube divides to produce a stalk cell and a spermatogenous (body) cell (Fig. 1.43D).

The spermatogenous cell enlarges consi­derably and two blepharoplasts develop at its two opposite end just before fertilisation. The spermatogenous cell divides vertically to form two sperm cells (male gametes) (Fig. 1.43E). The sperms of Ginkgo are very much similar to that of Cycas, except they are smaller in size, slightly elongated and spirally arranged cilia are mostly confined to the apical region.

Development of Female Gamelophyte:

The female gametophyte of Ginkgo deve­lops from the functional megaspore that enlarges considerbly and is surrounded by a thick mem­brane (Fig. 1.44A).

The nucleus of the megaspore divides mitotically, unaccompanied by wall for­mation, forming a large number of free nuclei (as much as 8,000 free nuclei) (Fig. 1.44B). The nuclei are restricted to a thin film of cytoplasm at the periphery resulting into the formation of a large central vacuole (Fig. 1.44C).

Thereafter, the cell wall formation begins in a centripetal fashion from periphery inwards (Fig. 1.44D), as a result the vacuole is obliterated. The entire gametophyte becomes cellular and the tissue thus formed is called endosperm (Fig. 1.44E). The endosperm cells are haploid in nature, but some polyploid cells are also formed. The cells that contain 2-3 nuclei during wall-formation are transformed to polyploid cells.

The ultrastructural study of the female gametophyte of Ginkgo shows four different donations according to their food reserves.

(i) Lipid zone (outer 3—4 layers),

(ii) Starch proteiolipid zone with large vacuolated cells,

(iii) Starch zone consists of large vacuolate cells with peripheral cytoplasm,

(iv) Central zone comprising of less differen­tiated cells with very little reserve content.

Development of Archegonia:

Two to four cells at the micropylar end of the female gemetophyte function as arche­gonial initials (Fig. 1.44F, 1.45A). Each of these cells divides periclinally to form on outer small primary neck initial and a large central cell (Fig. 1.45B). The primary neck initial divides by two vertical walls at right angles to each other resul­ting in four neck cells arranged in one tier.

The nucleus of central cell cuts off an upper ephemeral ventral canal cell and a large egg cells (Fig. 1.45C). With regard to ventral canal cell, Ginkgo seems to be more primitive than Cycas where a ventral canal nucleus is present (Fig. 1.45D).

At this stage, the female gameto­phyte grows upward forming a beak between the two archegonia (Fig. 1.44F, 1.46). The beak extends upward touching the inner surface of the nucellar beak like a pole in a tent and is known as ‘tent pole’.

Like Cycas, Ginkgo is anemophilous i.e., wind-pollinated. At the time of dispersal of pollen grains, the central strand of elongated nucellar cells in the pre-pollinated ovule disor­ganises followed by the collapse of epidermis resulting into a narrow and deep pollen cham­ber and pollination drop. The viscous fluid oozes out through the microphyle in the form of pollination drop.

Some of the anemophilous pollen grains are caught in the pollination drop and are brought to the pollen chamber due to the drying-off the fluid. Pollination generally takes place in May. Pollen grains germinate in the pollen chamber, unlike Cycas where they germinate in the intermediary chamber.

The pollen tube comes out through the aperture. The pollen tube branches freely showing a proximal unbranched tube and a much branched haustorial ramification in between the intercellular cells.

The sperms along with the pollen tube cyto­plasm are released in the archegonial chamber by the rupture of the terminal part of pollen tube. The osmotically rich cytoplasm of pollen tube causes the rupture of neck cells, as a result motile sperms enter into the archegonia. The fertilisation takes place in September i.e., four months after pollination.

The two sperms pro­ceed towards the archegonium with a forward and circular motion, ciliary band forming the posterior end. However, in Cycas, the ciliary band forms the anterior end. The archegonial chamber is filled up with the fertilisation fluid produced by nucellar cells. The ciliary band of the sperm is left behind on the top of the egg cell. The sperm nucleus fuses with the egg nucleus resulting in a zygote.

The zygote enlarges considerably and undergoes free nuclear divisions giving rise to 256 nuclei, evenly placed in the cytoplasm of the developing proembryo (Fig. 1.47A). The cell wall formations in the proembryo begin in all the cells simultaneously resulting in a cellular proembryo (Fig. 1.47B). The differentiation of proembryonal cells begins at a later stage.

The cells at the micropylar end elongate greatly forming a massive suspensor, while the basal cells (chalazal end) develop into embryonal cells with distinct meristem (Fig. 1.47C). A shoot apex is differentiate at the basal region of the embryo, while a root apex develops at the opposite end. A dicotyledonous embryo (rarely with three cotyledons) with two mesarch bundles is seen in a mature embryo (Fig. 1.48).

The most striking feature in Ginkgo is the development of embryo which continues even after the dispersal of seeds. Actually, the major part of the growth of embryo takes place in the detached seeds that are shed before or just after fertilisation. About seven months are required for attaining the maturity of an embryo.

Thus, Ginkgo shows the two-years reproductive cycle where the pollination takes place in spring, fertilisation in early autumn of the same year and the develop­ment of embryo continues even after shedding of seeds. The germination of the seed is hypogeal.


Yeast Strains, Culture, and Drugs

Yeast culture, including size analysis with a Coulter particle analyzer, fluorescence-activated cell sorting (FACS) analysis, and cell size selection by centrifugal elutriation has been described (Futcher, 1999 Jorgensen et al., 2002). Yeast strains are listed in Table 1. Alleles of bud21::BUD21 YFP -his5+ and sec63::SEC63 CFP -kanR were generated by genomic integration of standard C-terminal tagging cassettes that were PCR-amplified with integration site-specific primers. Template plasmids for cyan and yellow fluorescent protein (CFP and YFP, respectively) fusions were obtained from the Yeast Resource Center (University of Washington, Seattle). whi and CLN3 alleles have been described previously (Jorgensen et al., 2002). Synthetic complete medium is 0.2% amino acid mix, 0.17% yeast nitrogen base without amino acids and ammonium sulfate, 0.5% ammonium sulfate supplemented with 3% ethanol, 2% glucose, 2% raffinose, or 2% galactose, as indicated. YEP medium is 2% peptone, 1% yeast extract supplemented with 3% ethanol or 2% glucose, as indicated. Rapamycin and leptomycin B (LMB) were obtained from Sigma and were used at working concentrations of 0.2 μg/ml and 100 ng/ml, respectively.

Table 1. Strains used in this study

Microscopy and Morphometry

Sec63 CFP Reporter.

All experiments in Figures 1 –3 and 6 were performed with log phase cells at 30°C with OD600 <0.5 and cell concentration <3 × 10 7 cells/ml to prevent inadvertent nutrient depletion. Log phase cultures in synthetic media were rapidly concentrated by centrifugation. Medium, 2.5 μl, with concentrated live cells was mounted at room temperature and immediately visualized. Cell fields were imaged within 5 min of being removed from the 30°C incubator.

For Cells in Figures 1, 2, and 3, A and B.

Each cell field was sequentially imaged by differential interference contrast (DIC) and epifluorescent microscopy with an Eclipse E600FN microscope (100× objective, Plan Apo, Nikon, Melville, NY) and an Orca II CCD camera (Hamamatsu, Bridgewater, NJ). Sec63 CFP signal delineated the nuclear and cell envelopes. Metamorph (MDS Analytical Technologies, St. Laurent, QC, Canada) tools were used to individually outline nuclear, cell, and bud cross-sectional areas.

For Cells in Figures 3C and 6.

Each cell field was imaged as a small z-series, with five steps separated by 0.4 μm each. Sec63 CFP signal delineated the nuclear and cell envelopes. For each nucleus and cell, the largest cross-sectional area was determined from the z-series in Photoshop (Adobe Systems, San Jose, CA) and the cells and nuclei were converted to two separate binary masks that were analyzed in ImageJ ( using the “Particles 8 Plus” plugin ( in the Morphology package developed by Gabriel Landini (

For Cells in Figures 1 –3 and 6.

All cells in a field were quantified unless they were undergoing anaphase or cytokinesis, that is, if they possessed an elongated nucleus or two separated nuclei. For all experiments, multiple cell fields were imaged. The resulting data were analyzed in Microsoft Excel (Redmond, CA) and in Matlab (MathWorks, Natick, MA). For volume estimates, it was assumed that nuclei and G1 cells were spherical and that measured areas were cross-sections through the centers of these spheres.

NLS GFP Reporter.

Wild-type yeast (N-419) carrying the plasmid pMGGLA (ARS/CEN, LEU2, MET3pr-GFP-NLS-A) were cultured in synthetic -Met-Leu-Trp 3% ethanol medium. pMGGLA has been described previously (Edgington and Futcher, 2001). To enrich for small cells, the culture was elutriated, and 12 fractions of incrementally larger cells were obtained and kept on ice. Cells in fractions 1, 3, 4, and 10 were mounted. Slides were prepared with a small amount of molten agar mixed with minimal medium lacking a carbon source in microwells. Cells, 1 μl, were spotted onto the cooled, solidified medium, and a coverslip was applied. The coverslip edges were sealed with nail polish, the cells were examined with a microscope (Olympus) and photographed with a 1.3 megapixel Axiocam camera (Zeiss, Thornwood, NY), and morphometry was executed with Openlab software v.2 (Improvision, Lexington, MA). The perimeter of the nucleus was delineated by the edge of the green fluorescent protein (GFP) fluorescent signal, and the cell edge was obtained from DIC images.

Electron Microscopy

To generate a range of cell sizes, cells were collected from three different sources. All cells were in the W303 genetic background, whereas in the prior two approaches, all strains were from the S288c background. To measure small G1-phase cells, unbudded cells from a log phase, YEP 3% ethanol wild-type W303 culture were obtained by elutriation. Fifty-seven cell fields were analyzed by electron microscopy (EM) from these ethanol fractions. To measure moderately sized G1-phase cells, unbudded cells from a log phase, YEP 2% glucose wild-type W303 culture were obtained by elutriation. Twelve cell fields were analyzed by EM from these glucose fractions. To measure large G1-phase cells, conditional G1-cyclin depletion was performed by first propagating BS100 (cln1::LEU2-GAL-CLN1-HA3 cln2Δ::TRP1 cln3Δ::HIS3) to log phase in YEP 1% raffinose, 1% galactose. Cells were pelleted, washed twice in YEP without sugar, and then resuspended in YEP 2% glucose for 5 h. From these experiments, 16 BS100 cell fields were analyzed by EM. All samples were processed and photographed by Tamara Howard, a technician in Dr. David Spector's laboratory, Cold Spring Harbor Laboratory. Photomicrographs were scanned and analyzed using Image Reader 1.0 software from FujiFilm Science Lab for Windows (Stamford, CT). In each field of 30–50 cells, the cell with the largest cross-sectional nuclear area was chosen for further analysis, compensating for the fact that most sections were not taken through the center of the nucleus. It should be noted that our fluorescent studies indicate that there is not a strong relationship between nuclear size variability and cell size, so this method should not generate artifactual correlations between nuclear and cell size for that reason. For the chosen cell, nuclear and cell cross-sectional areas were measured.

Lobed Nuclei still count as One nucleus? - Biology

Blood provides a mechanism by which nutrients, gases, and wastes can be transported throughout the body. It consists of a number of cells suspended in a fluid medium known as plasma. Serum refers to plasma after clotting factors and fibrin have been removed.

Peripheral Blood Smear

The cells of the blood are important because they are a readily accessible population whose morphology, biochemistry, and ecology may give indications of a patient's general state or clues to the diagnosis of disease. For this reason, the complete blood count (CBC) and the differential white cell count are routinely used in clinical medicine. It is very important to be able to recognize normal blood cells and to distinguish pathological cells from the normal variants.

The identification of blood elements is based primarily on observations of the presence or absence of a nucleus and cytoplasmic granules. Other helpful features are cell size, nuclear size and shape, chromatin appearance, and cytoplasmic staining. The chart at the end of this section explains what to look for in the effort to identify the component cells of a blood smear.

Component Cell of the Blood Smear

A blood smear is created by placing a drop of blood near the end of a clean glass microscope slide. Another slide is held at an angle, backed into the drop, and then smoothly dragged forward to spread the blood film along the slide. The blood must then be fixed, stained, and washed.

When you view a properly prepared blood smear of a healthy individual, there are several populations of cells that you will notice. Keep in mind that these are all mature cells. The next section will discuss the identification of the immature cells of the bone marrow.

  • Erythrocytes, or red blood cells, are by far the predominant cell type in the blood smear. They are anucleate, non-granulated, eosinophilic cells that are uniform in shape (biconcave discs) and size (7.2 microns). Red blood cells have a central concavity that appears pale under the light microscope. These cells contain hemoglobin and are responsible for the transport and delivery of oxygen. Erythrocytes have a lifespan of 120 days.
  • Reticulocytes are immature red blood cells that are released from the bone marrow. They mature into erythrocytes after 1 to 2 days in the peripheral blood. There should be about one reticulocyte for every 100 red blood cells in a normal blood smear. These cells stain with a light blue tint because they still have RNA-containing organelles like free ribosomes.
  • Thrombocytes, or platelets, are the smallest elements of the blood and are responsible for the formation of clots through a complex, highly regulated cascade that you will study in Physiology and Immunobiology. Platelets are between 2 and 5 microns in diameter and appear ovoid and anucleate with purple granules.
  • Leukocytes, or white blood cells, are cells of the immune system that are present in both blood and interstitial fluid. There should be about 1 leukocyte for every 1000 red blood cells. They can be classified into two groups according to their nuclear pattern and the presence of cytoplasmic granules.

Monomorphonuclear leukocytes are cells with round, non-lobed nuclei These include:

  • Small lymphocytes, which are about the same size as erythrocytes and have deeply stained nuclei with a thin rim of cytoplasm. This population includes both B-cells and T-cells.
  • Large lymphocytes, which appear similar to small lymphocytes, but with larger nuclei and a greater amount of cytoplasm. This population also includes both B-cells and T-cells. Lymphocyte counts are raised in response to viral infections.
  • Monocytes, which are larger than lymphocytes and have less-clearly demarcated nuclei that are usually not centered in the cell. These nuclei appear horseshoe-shaped and the cytoplasm contains fine granules that give it a muddy gray color. These granules contain lysosomal enzyme and peroxidase. Monocytes are phagocytic cells that are important in the inflammatory response. They are the precursors to the tissue macrophages that you studied in the Laboratory on Connective Tissue.

Polymorphonuclear leukocytes are cells with lobed nuclei and cytoplasmic granules. While these cells share the same primary (nonspecific) or azurophilic granules, they are named based upon the characteristics of their secondary (specific) granules.

  • Neutrophils are by far the most numerous of the leukocytes. They are characterized by a nucleus that is segmented into three to five lobes that are joined by slender strands. The cytoplasm of neutrophils stains a pale pink. Its primary granules contain acid hydrolases and cationic proteins, and its secondary granules contain a variety of antimicrobial substances used to destroy bacteria that they phagocytose during the acute inflammatory response.
  • Eosinophils are larger than neutrophils and are distinguished by large red or orange granules of uniform size. These granules contain major basic protein, which is released to kill organisms too large to phagocytose, such as parasites and helminthes (worms).
  • Basophils are intermediate in size between neutrophils and eosinophils and have simple or bilobed nuclei. They contain many coarse purple granules that can vary in size or shape. These granules contain histamine, which is released to cause a vasoactive response in hypersensitivy reactions, and heparin, which is an anticoagulant. Basophils are not phagocytic.

Component Cells of a Bone Marrow Smear

While the peripheral blood smear indicates the status of mature blood cells, the bone marrow smear can be used to assess the process of hematopoiesis, or blood cell formation.

Active bone marrow appears highly cellular. The majority of the developing cells will become erythrocytes, which confer a red color to the marrow. For this reason, active bone marrow is also known as red bone marrow. Over time, the marrow becomes less active and its fat content increases. It is then referred to as yellow bone marrow.

Once again, there are several important characteristics to take into account when viewing a bone marrow smear. These include:

  • Size of the cell
  • Cytoplasm to nucleus volume ratio
  • Shape of the nucleus
  • Degree of chromatin condensation
  • Presence or absence of nucleoli
  • Cytoplasmic staining
  • Presence of cytoplasmic granules

The blast cell is a pluripotent stem cell from which erythrocytes, granulocytes, and lymphocytes originate. Erythrocytes develop from erythryoblasts, granulocytes from myeloblasts, and lymphocytes from lymphoblasts. These cells, however, all appear identical - they are large with round or ovoid nuclei, a distinct nuclear membrane, visible nucleoli, and an abundant blue cytoplasm. As the blast cells differentiate, the resultant cells can be assigned to a particular cell line.

Erythropoiesis is the development of red blood cells. There are several recognizable steps in this lineage:

  • The erythroblast develops into a proerythroblast, which is only slightly smaller than the blast, but has a more basophilic cytoplasm.
  • The basophilic erythroblast forms when the proerythroblast loses its nucleolus. These cells are much smaller than the blast cells and have an intensely basophilic cytoplasm that results from the accumulation of ribosomes.
  • The polychromatophilic erythroblast has a darkly staining nucleus and its cytoplasm stains a grayish-green color due to the accumulation of hemoglobin.
  • In the orthochromatic erythroblast, or normoblast, the nucleus becomes smaller and darker and the cytoplasm becomes pinker. Nuclear expulsion occurs at the end of this stage through an asymmetric division of the orthochromatic erythroblast. The portion that contains the cytoplasm and organelles becomes the reticulocyte, while the portion containing the nucleus is destroyed by macrophages.
  • The reticulocyte contains cytoplasm, cytoplasmic organelles, and many ribosomes. It is released from the bone marrow and develops into a mature erythrocyte after spending 1 to 2 days in the peripheral blood.

Granulopoiesis is the process by which white blood cells develop. The myeloid series has the most characteristic cell lineage:

  • The myeloblast differentiates into a promyelocyte that becomes irreversibly committed to the neutrophilic cell line. This cell is large, with a large round nucleus, prominent nucleoli, and purple azurophilic granules. These granules are primary, nonspecific granules. Promyelocytes also give rise to eosinophils and basophils
  • The myelocyte stage is characterized by the production of secondary, specific granules. Myelocytes can vary in cell size and nuclear shape. They contain both the purple staining azurophilic granules and lilac staining specific granules. As they develop, they decrease in size, their nucleus becomes indented, and there is a shift toward more specific granules. There is also a reduction in the number of organelles, which results in decreased basophilia of the cytoplasm.
  • The metamyelocyte has a flattened nucleus with condense chromatin.
  • The band cell has a horseshoe-shaped nucleus that is "immature." As development continues, it will mature into a segmented nucleus with multiple lobes. It will then be a mature neutrophil.

Eosinophils and basophils undergo sequential stages of differentiation in a very similar manner to those of neutrophils. Their specific granules are also produced during the myelocyte stage.

The platelet lineage is similar. Large, multilobed promegakaryocytes develop into megakaryocytes, which are the largest cells of the bone marrow (30 to 40 microns). Platelets form through the segmentation of these cells.

Monocytes develop from promonocytes and lymphocytes develop from prolymphocytes. These elements are difficult to distinguish in normal bone marrow smears.

Life Cycle of Penicillium (With Diagram) | Fungi

The mycelium is well developed and copiously branched. It is composed of colourless, slender, tubular, branched and septate hyphae. The hyphae run in all directions on the substratum and become intertwined with one another to form a loose network of hyphae constituting the mycelium.

Some of the hyphae may even grow into the interior of the substratum and the rest spread on the surface. The former secrete enzymes and absorb food materials from the substratum. These are the haustorlal hyphae.

The aerial hyphae receive nourishment through the haustorial hyphae and produce reproductive structures. Baker (1944) observed anastomosing between hyphae of two mycelia resulting in a heterokaryotic mycelium.

The mycelium in a few species may develop into a sclerotium. The hyphae constituting the mycelium are septate and the cells are short. The septa between the cells have each a central pore. Through the pores the protoplasm flows from cell to cell.

Reproduction in Penicillium:

Penicillium reproduces both asexually and sexually. The asexual stage however, is dominant and constitutes the usual mode of reproduction. Sexual stage is rare.

1. Asexual Reproduction:

It takes place by vegetative methods and sporulation.

(i) Vegetative Reproduction:

It is accomplished by the most common method of fragmentation. The hyphae break up into short segments. Each segment or fragment grows by repeated division into a full-fledged mycelium.

In some species, the mycelium forms compact resting bodies, the sclerotia. Sclerotia enable the species to survive periods of stress or to hibernate. On the onset of conditions favourable for growth each sclerotium germinates to form a new mycelium. The sclerotia thus serve primarily as a means of perennation rather than multiplication.

Normally it takes place by the formation of non-motile, asexual spores, the conidia which are produced exogenously at the tips of long, erect special septate hyphae called the conidiophores. Penicillium multiplies repeatedly by this method during the growing season.

Conidiophores (Fig. 10.10 A-C):

A conidiophore arises as an erect, tubular hyphal outgrowth from any cell of the mycelium and not from a specialised cell (foot cell) as in Aspergillus. After some period of vegetative growth upright hyphae arise from the older portions of the mycelium.

They are negatively geotropic and arise singly from any cell of the mycelium. Each grows up in length vertically. Reaching a certain height the septate conidiophore branches once or twice or even more times.

These are termed as primary, secondary or tertiary branches, respectively. Only rarely are the conidiophores unbranched (Penicillium thomii). The unbranched axis of the latter bears a tuft of flask-shaped sterigmata (A).

In the species with branched conidiophores, the ultimate branches which bear tufts of flask-shaped sterigmata or the phialides are called the metulae. The lower branches which support the metulae when short and form a part of the penicillus are called the rami.

Conidia are abstricted from the tips of the phialides or sterigmata and are borne in long, unbranched chains. Baker (1944) reported that the phialides and the upper cells of the conidiophore are uninucleate.

The phialide wall is electron-transparent with a thin electron opaque surface layer. The apical portion of the conidiophore with its branches (metulae), sterigmata and chains of conidia looks like a small artist’s brush known as the penicillus.

The generic name Penicillium is derived from ‘penicillus,’ which reflects the form of conidial chains arising from the sterigmata supported on the metulae.

Development of Conidia:

The conidia are formed within the narrow tips of the flask-shaped phialides (A). The conidium initial is formed by the distension of the tubular tip of the phialide (B). The phialide nucleus undergoes mitosis. One daughter nucleus remains in the phialide and the other migrates into the swollen tip (conidium initial). The conidium initial protoplast is then cut off from the phialide protoplast by a thin perforate septum (C).

The perforation remains as a channel between the successive conidia in the chain. It is filled with electron-opaque material. The newly delimited conidial protoplast secretes a wall around it distinct from the phialide wall and functions as the first conidium (D).

The spore or conidial wall during further development may remain distinct or fuse partially with phialide wall. The tip of phialide below the first conidium again elongates and swells (D). A second conidium chains formed by repeating the process (E).

Like this, one below the other, a long chain of conidia is formed. In older conidium chains, the septum region remains only as a narrow, connecting strand between conidia. The conidium chains are enclosed within an electron-opaque surface layer which appears to be continuous with the surface layer of the phialide wall.

The conidium in the chain are arranged in a basigenous manner. The youngest conidium lies next to the tip of the sterigma and the oldest away from it.

The basigenous arrangement of conidia serves two useful purposes. It permits ready dispersal of mature conidia from the tips. Secondly, it aids the proper nourishment of younger conidia which are nearest to the tips of the sterigmata.

As the conidial chain increases in length, the connectives between the older conidia break down resulting in the separation of mature conidia. The conidia are thus shed continuously. Being small, light and dry they are dispered by air currents.

The conidia under the microscope look like small beads. They may be ovoid globose, elliptical or pyriform. In some species they are smooth and in others rough. Generally the conidia are uninucleate but may become multinucleate in some species.

The conidia may be greenish or pale in colour depending upon the species. It is the spore wall that is coloured. The conidia are responsible for the colony colour characteristic of the species. They serve for quick propagation of the species during the growing season. They are easily disseminated by wind.

The conidia are tiny spore-like structures globose to ovoid in form. The pigmented spore wall is differentiated into two layers, outer exine and inner intine. The exine is comparatively thick, smooth or spiny. The inline is thin. Under electron microscope it appears to consist of 3 or 4 layers. There is the outermost layer (W1) with an irregular, undulating contour. It is electron-dense.

Empty speaces are often seen between it and the next wall layer. Next to it is a slightly thin inner electron-opaque layer W2 with a more regular contour. This layer (W2) gradually merges into the next wall layer W3 which is thick and electro- transparent. It usually abuts on the plasma membrane. However, in some cases, there is another slightly electron-dense layer (W3) present directly above the plasma membrane. Within the spore wall is the plasma membrane.

Embedded in the cytoplasm of the conidium are the mitochondria and ribosomes. The endoplasmic reticulum strands are not discernible. The vacuoles are absent. However, Martin et al. (1973) reported the presence of a single large vacuole in the resting conidium of P. notatum. The conidium cytoplasm contains oil globules. Usually the conidium contains a single nucleus. The nuclear membrane is two-layered and is poriferous.

The conidial stage in Penicilliutn is more dominant than in Aspergillus. Majority of the species of Penicillium are known only in the conidial stage. Some (about 20 species) are now known to produce cleistothecia.

At first when the connection between the two stages was not fully established the conidial or imperfect stage was given the name form or genus Penicillium and the sexual or perfect stage as Talaromyces. The discovery of sexual or perfect stage in these form- species places them in the true Ascomycete genus Talaromyces.

Many mycologists, however, still adhere to the old name Penicillium because the conidial stage is predominant. Moreover, the generic name Penicillium was introduced first.

Germination of Conidia (Fig. 10.13):

On falling on a suitable substratum and under suitable conditions (moisture, suitable temperature and food), the conidium absorbs moisture and swells. The swollen conidium germinates by putting out a germ tube. Fletcher (1971) studied the fine structural changes during germination of conidia in Penicillium griseofulvum. He reported that the ungerminated conidium has a two-layered spore wall.

The protoplast contains a single nucleus and mitochondria. During germination a third layer appears inside the two-layered spore wall. It is continuous with the germ tube wall. In P. frequentans, the germ tube wall is considered to be an extension of the inner layer of the original spore wall.

During swelling the mitochondria increase in size and become lobed, endoplasmic reticulum becomes visible md vacuoles are formed. The septa formed in the germ tube are perforate and have Woronin bodies associated with them. Marchant (1968) suggested enzymic breakdown of the two-layered spore wall at the point of germ tube emergence rather than mechanical rupture.

Martin et al. (1973) reported that during swelling of the conidium (A), the outer layer (W1) of the wall of the conidium becomes broken at several places and separated from the underlying spore wall layer W2 . The smooth surface layer W2 with a regular contour thus becomes exposed. At this stage the swollen conidium pats out a germ tube (B).

During spore swelling the large mitochondria of the resting spore divide to produce smaller mitochondria. The size of the single vacuole decreases. Endoplasmic reticulum reappears in the swollen and germinating spores. The wall of the emerging germ tube has been reported to be continuous with the electron transparent wall layer W3.

The electron-dense wall layer W2 remains at the base of the germ tube and around the spore. The single nucleus of the resting spore divides early in germination. One of these migrates into the germ tube.

With the elongation of the germ tube most of the mitochondria migrate into it and become concentrated near the tip. Small vesicles reappear along the plasma membrane. Some were seen at the tip of the germ tube. Later, a septum is formed at the point of emergence of the germ tube. It is continuous with the transparent layer W3 of the spore wall (C).

During further development, layers W2 and W1 of the hyphal wall are formed by deposition of electron-dense material on the outer surface of the germ tube. The latter elongates and divides by septa. Septum formation is preceded by nuclear division. Each septum has a central pore.

There are Woronin bodies associated with the septa. By further growth, septation and branching mycelium is formed. It consists of uninucleated cells.

The mycelial hyphae of Penicillium, when made to grow immersed in a sugary solution, they divide by additional septa into short uninucleate segments. The latter become rounded and separates as thin-walled spore-like structures called the oidia or oidiospores.

Often the oidia increase in numbers by budding. This is known as the torula stage. The fungus m the torula condition brings about fermentation of sugar into alcohol. This process is called alcoholic fermentation. On a solid medium each oidium germinates to produce a normal mycelium.

Similarly if conidia of Penicillium happen to fall in a sugary solution they fail to produce normal mycelia. Instead each conidium germinates to produce a thin-walled yeast-like cell which starts budding like the oidiospore to produce the torula stage.

2. Sexual Reproduction (Fig. 10.14):

Sexual process in Penicillium has been studied in a few species such as P.vermiculatum (=Talaromyces vermiculatus), P. glaucum, P. brefeldianum and a few others. All of them are reported to be homothailic. Derx (1925) reported heterothallism in one species (P. luteum) but his observation has not been confirmed by subsequent investigators.

The structure of sex organs varies from species to species. In Talaromyces vermiculatus (Penicillium vermiculatum) sexual reproduction is oogamous. It was studied in detail by Dangeard in 1907. The male and female organs are known as antheridia and ascogonia respectively.

The mature ascogonium is a long, erect multinucleate, unseptate, tubular structure. At its upper end it may be curved like the handle of an umbrella (E). It arises as a lateral outgrowth from any cell of the vegetative mycelium. When young the ascogonium is uninucleate (A). As it elongates the single nucleus within it divides and redivides to give rise to a definite number of daughter nuclei which is either 32 or 64 (B).

Meanwhile a slender uninucleate hyphal branch originates either from an adjacent cell of the same hypha which gives rise to the ascogonium or from a separate neighbouring hypha. It is the antheridial branch (C). It grows up coiling loosely around the ascogonium making several turns about it (D).

The distal end of the male branch becomes slightly inflated and is eventually cut off as an antheridium by a septum (E). The antheridium is a short, terminal, club-shaped, uninucleate structure.

It is the union of two protoplasts which brings the compatible nuclei close together in the same cell. The tip of the antheridium comes in contact with the ascogonium. At the point of contact, the double wall dissolves. Through the common pore the two protoplasts come in contact. According to Dangeard, the migration of the male nucleus into the ascogonium, does not take place.

This fact has been confirmed by a number of other workers. Mere contact of the antheridium protoplast stimulates the ascogonium. Beyond that the antheridium plays no role. The female nuclei in the ascogonium, however, arrange themselves in pairs.

The pairing of the female nuclei in the ascogonium is called autogamy. Each pair is called a dikaryon. With the establishment of dikaryons the haplophase ends and the dikaryophase starts in the life cycle.

In some other species of Penicillium, both antheridium and ascogonia are claimed to be functional. The intervening walls between the two sex organs dissolve and their protoplasts come in contact. The male nucleus migrates into the ascogonium. Further details of the male nucleus are not known fully.

The suggestion, however, is that it divides a number of times. The male nuclei then come to lie by the side of the female nuclei, each to each. Each pair of nuclei consisting of one male and the other female is called a dikaryon.

Sexual reproduction has been worked out in P. glaucum. Two short lateral hyphae arise from the mycelium close together and become coiled about each other. One of them is the antheridium and the other ascogonium. Plasmogamy thus takes place by gametangial contact. Dikaryons are established in the ascogonium. This is followed as usual by the septation of the ascogonium into binucleate segments.

Post Plasmogamy or Autogamy changes (Fig. 10.14 E-G):

(i) Septation of Ascogonium:

Plasmogamy or Autogamy is followed by septation of the ascogonium. Each segment has a pair of nuclei. Meanwhile entangled sterile hyphae grow up around the sexual apparatus and form an investment of loosely interwoven hyphae and sexual apparatus and afford protection to the structures developing within (E-F).

(ii) Development of Ascogenous Hyphae (Fig. 10.14 F):

As a result of the stimulus of plasmogamy or autogamy one or more lateral outgrowths arise from some of the binucleate segments situated in the middle of the septate ascogonium. Each outgrowth is called an ascogenous initial.

It is binucleate. Each ascogenous initial develops into a branched ascogenous hypha composed of binucleate cells. The branches are of different lengths. In some other species of Penicillium the lateral branches of ascogenous hyphae are one-celled.

(iii) Formation of Asci:

According to Emmons (1935), asci in many species are developed directly from the binucleate cells toward the tips of the branches of the ascogenous hyphae. Consequently they are arranged in short chains. In P. vermiculatum almost all the cells of the ascogenous hyphae develop into asci directly without crozier formation. Rarely the asci are formed from the croziers.

1. Direct Method of Ascus Development:

This is illustrated by P. vermiculatum (Fig. 10.15). All the binucleate cells of the ascogenous hyphae are capable of directly developing into ascus mother cells. No croziers are formed. The binucleate cell enlarges into a sac-like structure. It is the ascus mother cell.

It is globose or pear-shaped in form. The two nuclei of the ascus mother cell eventually fuse. The cell containing the fusion nucleus (synkaryon) is called the young ascus.

2. Indirect (crozier) method of ascus formation (Fig. 10.16):

In this case, the terminal binucleate cells of the ascogenous hyphae or their branches curl over to form a hook-like structure called the crozier. For further details of the process refer to prior pages.

The binucleate cell in both cases enlarges to function as the ascus mother cell. It is the last structure of the dikaryophase. The two nuclei in it eventually fuse. This is karyogamy. Karyogamy is equivalent to fertilisation. With karyogamy, the dikaryophase in the life cycle ends and the diplophase begins.

The septate ascogonium, ascogenous hyphae fusion or diploid nucleus (synkaryon) is the young ascus. It is the only diploid structure in the life cycle and thus represents the short-lived, transitory diplophase.

(iv) Differentiation of Ascospores Fig. 10.16 (F-I):

The young ascus enlarges. Its— diploid nucleus undergoes three successive divisions. The first and the second divisions constitute meiosis. The third is mitotic. The resultant 8 nuclei are thus haploid. A small amount of cytoplasm gathers around each daughter nucleus.

The uninucleate protoplasts secrete their own walls and are fashioned into uninucleate ascospores. Eight ascospores are thus formed by the method of free cell formation in each mature ascus which may be spherical or pear-shaped in appearance.

(v) Formation of Ascocarp (Fig. 10.17):

With the septation of ascogonium and development of ascogenous hyphae, a large number of sterile hyphae grow up around the sexual apparatus. The ensheathing sterile hyphae get interwoven to form a hollow ball- like structure, the peridium which surrounds and protects the ascogenous hyphae as they grow and branch within.

This is the ascocarp. The asci within the ascocarp are scattered. The ascocarp of Talaromyces is of indefinite growth. It continues to increase in size even after the after the ascospores begin to mature.

The peridium or sheath is thicker than that of the ascocarp of Aspergillus and consists of loosely interwoven hyphae. This entire structure is spherical and has no opening (A). Such a closed fruit or ascocarp is called the cleistothecium. The asci are borne in chains within the ascocarp.

In some other sp. of Penicillium, the peridium is compact and pseudoparenchymatous. The asci are borne singly and terminally on one-celled lateral branches of the ascogenous hyphae within it.

The cleistothecium of Penicillium or Aspergillus represents the following three generations:

It is represented by the sheath or peridium made up of loosely interwoven hyphae.

It consists of the binucleate cells of the ascogonium, ascogenous hyphae and the ascus mother cells. The transitory diplophase is represented by the young ascus containing a diploid nucleus.

(iii) Future haplophase:

It is represented by the ascospores in the asci.

Discharge of Ascospores:

At maturity the walls of the asci dissolve. The liberated ascospores, each shaped like a pulley wheel (side view), float in the nourishing fluids, formed by the degeneration of the inner layers of the peridium, walls of asci and the ascogenous hyphae. The ascospores absorb nutrition and mature. The mature ascospores are finally released by the decay of the other wall of the peridium.

Ascospore Structure (Fig. 10.18):

The liberated ascospore is a haploid uninucleate structure. It is lens-shaped with a small groove around the edge and thus appears like a pulley wheel in side view (B). The spore wall may be smooth or sculptured. It is differentiated into two layers, the outer epispore and inner endospore. The epispore is usually thick and sculptured. In face view the ascospore is round to star-shaped (A).

Alteration of Generations in Penicillium Brefeldianum:

In this species the sexual apparatus (antheridia and ascogonia) is absent. The vegetative hyphae give rise to two similar protuberances. They are known as the copulation branches.-The two copulation branches coil around each other. Their tips come in contact. At the point of contact the walls between the tips dissolve. The contents of one migrate into the other to establish a dikaryon in the fusion cell.

This species of Penicillium provides an example of isogamous sexual reproduction. After the migration of the contents the copulation branches are surrounded by sterile hyphae. The latter arise from the adjacent cells of vegetative hyphae. These sterile hyphae form a hyphal knot. The latter subsequently assumes a plectenchymatous character.

After about a week the outer membrane of the hyphal knot thickens to form a hard outer covering. The fusion or dikaryotic cell, in the meantime, sends out certain outgrowths. They are unseptate and are known as the ascogenous hyphae.

The dikaryon in the dikaryotic cell undergoes conjugate division. A pair of nuclei passes into each ascogenous hypha. The ascogenous hyphae become divided into short and cylindrical binucleate cells. Some of the cells grow out into buds with spirally coiled tips.

From the convex surface of these spirally coiled tips there arise branches that curve upwards and coil. From these branches there again arise short multicellular branches. Each cell of these last formed branches becomes spherical. It is known as an ascus. Nothing is known about the nuclear fusions in the ascus. Each ascus contains eight ascospores.

The asci containing ascospores, the ascogenous hyphae and the surrounding sterile hyphae with their outer hardened rind constitute a fructification. It is closed and is called the cleistothecium. The walls of the asci degenerate and the ascospores are liberated in the centre of the cleistothecium.

Later the outer hard rind of the cleistothecium also breaks. As a result the ascospores are released. Each ascospore possesses a double membrane the outer exospore and the inner endospore.

Life Cycle of Vaucheria (With Diagram) | Xanthophyta

The vegetative reproduction takes place by fragmentation. The thallus can break into small fragments due to mechanical injury or insect bites etc. A septum develops at the place of breaking to seal the injury. The broken fragment develops thick wall and later on develops into Vaucheria thallus.

2. Asexual Reproduction in Vaucheria:

The asexual reproduction takes place by formation of zoospores, aplanospores and akinetes

The zoospores formation is the most common method of reproduction in aquatic species. In terrestrial species it takes place when the plants are flooded. Zoospore formation takes place in favourable seasons or can be induced if aquatic species are transferred from light to darkness or from running water to still water.

Zoospores are formed singly within elongated club shaped zoosporangium (Fig. 2A, B). The development of zoosporangium begins with a club shaped swelling at the tip of a side branch. A large number of nuclei and chloroplasts along with the cytoplasm move into it. A colourless protoplasmic region becomes visible at the base of cytoplasm and it is separated from rest of the cytoplasm of thallus.

Each separated protoplast secretes thin membrane and zoosporangium gets separated by a cross wall. Inside zoosporangium the vacuole decreases, the contents of sporangium become very dense and round off. The change takes place in relative position of chloroplasts and nuclei, the nuclei become peripheral and chloroplasts enter in inner layer of cytoplasm.

The entire protoplasm of the zoosporangium contracts to form oval zoospore. Opposite to each nucleus two flagella are produced making zoospore a multi-flagellate structure. A terminal aperture develops in zoosporangium by gelatinization of wall. The zoospore is liberated through aperture in morning hours (Fig. 2 C, D).

Each zoospore is large yellow green, oval structure. It has a central vacuole which has cell sap and may be traversed by cytoplasmic strands. The protoplasm outer to vacuole has many nuclei towards the walls and chromatophores towards vacuoles. Two flagella arise opposite to each nucleus. This part of cytoplasm can be regarded equivalent to one zoospore.

Fritsch (1948) regarded this kind of zoospore as compound zoospore or synzoospore as a number of biflagellate zoospores have failed to separate from one another.

According to Greenwood, Manton and Clarke (1957) the flagella of a pair are heterokontic and whiplash type. The shorter flagellum of each pair is directed towards the anterior end of the zoospore. The flagellar bases are united together in pairs and are firmly attached to the tip of nuclei.

According to Greenwood (1957), there is large anterior vacuole and small ones in the posterior region of the zoospores. Mitochondria are present in the peripheral layer of cytoplasm. Fat bodies and plastids are present in the cytoplasm. Chlorophyll has also been reported from the zoospores.

The zoospores swim in water for 5-15 minutes and germinate without undergoing any significant period of rest. The zoospores get attached to the substratum, withdraw flagella and secrete thin walls (Fig. 2 E, F). The chromatophores move outwards and nuclei inwards as in vegetative condition.

The two tube like outgrowths develop in opposite directions. One of the two outgrowths elongates, branches to form colourless lobed holdfast and the other outgrowth forms yellow-green tubular coenocytic filament (Fig. 2 G, H).

Aplanospores are commonly observed in species. V. geminata, V. uncinata and in marine species V. pitoboloides. The aplanospores are generally formed by terrestrial species.

Aquatic species form aplanspores under unfavorable condition of drought. The aplanospores are non-motile asexual spores formed in special structures called aplanosporangia (Fig. 3 A-C). The aplanospores are produced singly in cells at the terminal end of the short lateral or terminal branch.

The protoplasm of aplanosporangium gets metamorphosed into single multinucleate aplanospore which is thin walled. In V. germinata aplanospores are oval and are liberated from apical pore formed by gelatinization.

In V. uncinata aplanospores are spherical and are liberated by rupture of the sporangial wall. The formation and structure of aplanospores and zoospores is similar except that the zoospores lack flagella. The aplanospores soon after liberation germinate into new thalli (Fig. 3D).

Akinetes are thick walled structures formed during unfavorable conditions like drought, and low temperature. The akinetes have been commonly observed in V. geminata, V. megaspora and V. uncinata.

The akinetes are formed on the terminal part of lateral branches where protoplasm migrates to the tips followed by cross-wall formation (Fig. 4). These multinucleate, thick walled segments are called akinetes or hypnospores.

The akinetes by successive divisions may form numerous thin walled bodies called cysts. When many akinetes remain attached to the parent thallus, the thallus gives the appearance of another alga Gongrosira.

Hence this stage of Vaucheria is called Gongrosira stage. During favourable conditions the akinetes and cysts develop into new thalli. Randhawa (1939) has reported that in V. uncinata the submerged parts of thallus develop sex organs whereas exposed parts of thallus form brick shaped akinetes.

(iii) Sexual Reproduction in Vaucheria:

In Vaucheria sexual reproduction is of advanced oogamous type. The male and female sex organs are antheridia and oogonia, respectively.

Majority of the freshwater species are monoecious or homothallic while some species like V dichotoma, V. litorea and V. mayyanadensis are dioecious or heterothallic. There are different types of arrangement of antheridia and oogonia in homothallic species. The position, structure and shape of antheridia are of taxonomic importance in Vaucheria.

The common patterns of arrangement of sex organs are as follows:

(a) Antheridia and oogonia develop close to each other on the filament at intervals (Fig. 5 A-C).

(b) The antheridia and oogonia are borne on special side branches with a terminal antheridium and a number of lateral oogonia (Fig. 5D).

In V. hamata the reproductive branches bear a median terminal antheridium and two oogonia, one on either side of antheridium.

In V. geminata and V. terrestris the sex organs are produced at the ends of the lateral branches with a terminal antheridium and a group of oogonia (Fig. 5D). The sex organs are unilateral when they are arranged on one side of the filament or bilateral when they are on both sides of the filament.

(c) Antheridia and oogonia are borne on adjacent branches (Fig. 5E).

Structure and Development of Antheridium:

The mature antheridia may be cylindrical, tubular, straight or strongly curved. The antheridium is separated from main filament by a septum. The antheridia can be sessile (without stalk) arising directly from main branch e.g., V. civersa. The antheridia may be placed high on the branch the antheridia are situated on androphore V. synandra.

The young antheridium is usually green in colour. It contains cytoplasm, nuclei and chloroplasts. The mature antheridia are yellow and contain many spindle shaped antherozoids. The antherozoids are liberated through a terminal pore e.g., V. aversa or through many pores e.g., V. debaryana

In monoecious species the antheridium arises as a small bulging or lateral outgrowth along with or before the oogonium development (Fig. 6A). Many nuclei along with cytoplasm enter into it and it gets cut off from the lower part forming a septum.

The antheridium grows and becomes high curved structured, its upper part is main antheridium and the lower part is stalk. The nuclei of antheridium get surrounded by cytoplasm and develop into biflagellate, yellow coloured antherozoids The antherozoids are liberated from the tip of antheridium through apical pore shortly before day break (Fig. 6D-1).

Structure and Development of Oogonium:

The oogonium development starts with accumulation of colourless multinucleate mass of cytoplasm near the base of antheridial branch. This accumulated cytoplasm has been termed as “wanderplasm”. The wanderplasm enters into the outgrowth or bulging of the main filament. This outgrowth is called as oogonial initial.

Large amount of cytoplasm and nuclei enter into oogonia, making it a large globular structure called as oogonium (Fig. 6 B-E). As the oogonium matures, it gets separated from main branch by the development of septum at its base. The mature oogonium is uninucleate structure. The nucleus of oogonium with protoplasm develops into a single egg.

There are three hypothesis regarding the fate of extra nuclei of oogonium of Vaucheria:

(a) According to Oltmanns (1895) accept a single nucleus which forms female nucleus, all other nuclei migrate back into the filament. This was supported by Heidinger (1908) and Couch (1932).

(b) According to Davis (1904), the single nucleus forms the egg and all other nuclei degenerate.

(c) According to Brehens (1890) all nuclei fuse to form a single nucleus.

The mature oogonia are globose, obovoid, hemispherical or pyriform in shape. The oogonia may be sessile or stalked structure. The protoplast of oogonium is separated from main filament by- septum formation.

The entire protoplasm with single nucleus makes a central spherical mass called as oosphere or ovum. In mature oogonium a distinct vertical or oblique beak develops in apical part. Opposite to beak develops a colourless receptive spot. A pore develops just opposite to receptive spot (Fig. 6 F).

The oogonium secretes a gelatinous drop through a pore near the beak. A large number of liberated antherozoids stick to the drop. Many antherozoids push into the oogonium. The antherozoids strike violently, fall back and push forward again and fall back. Only one antherozoid enters into the oogonium.

After its entry the membrane develops at the pore to stop the further entry of antherozoids. The male nucleus increases in size and fuses with the egg nucleus to make diploid zygote. The zygote secretes a thick 3-7 layered wall and is now called as oospore (Fig. 6 G-I). The chromatophores degenerate and lie in the centre of the cell.

Germination of oospore:

The oospore undergoes a period of rest before germination. During favourable season the oogonial wall disintegrates and the oospore is liberated. The oospore germinates directly into new filaments.

Although the exact stage at which the reduction division takes place in Vaucheria is not clear, it is believed that reduction division occurs in first nuclear division in the germinating oospore (Fig. 7 A-D). The oospore germinates to make haploid thallus of Vaucheria.

According to Williams, Hanatsche and Gross the life cycle of Vaucheria is haplontic, the oospore being the only diploid structure in life cycle (Figs. 8, 9). Vaucheria thallus is haploid. It is aseptate, branched, tubular and coenocytic structure.

Vegetative re-production takes place by fragmentation. Asexual reproduction takes place by zoospore in aquatic species and by aplanospores in terrestrial species.

The zoospore is large multi flagellate structure and is supposed to be compound:

Zoospore or Synzoospore:

The sexual reproduction is advanced oogoinous type, the male and female sex organs are antheridia and oogonia. Most of the species are homothallic, some are heterothallic. After fertilization, a diploid zygote is formed which converts into oospore and undergoes a period of res The reduction division takes place in oospore during germination and an haploid thallus is formed (Fig. 8, 9).

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